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Full text of "soil analysis"

Methods of 
Phosphorus Analysis 

for Soils, Sediments, Residuals, and Waters 



Southern Cooperative Series 

Bulletin No. 396 

A Publication of SERA-IEG 17 

A USDA-CSREES Regional Committee 

Minimizing Agricultural Phosphorus Losses lor 

Protection of Ihe Water Resource 






. 




AAESD, 



Southern Cooperative 
Series Bulletin 



Methods of Phosphorus Analysis for Soils, 
Sediments, Residuals, and Waters 

Southern Cooperative Series Bulletin No. # 396 



June, 2000 

URL 

http://www.soil.ncsu.edu/seral7/publications/seral7-2/pm_cover.htm 

Contact Information 



Gary M. Pierzynski Editor 



Department of Agronomy 
2004 Throckmorton Plant Sciences Ctr. 
Kansas State University 
Manhattan, KS 66506-5501 



ISBN: 1-58161-396-2 



This bulletin, from the joint Southern Extension/Research Activity - Information Exchange Group (SERA- 
IEG) 17, included extension specialists and research scientists. It is being published with the approval of 
the Southern Association of Agricultural Experiment Station Directors (SAAESD) and the Association of 
Southern Region Extension Directors (ASRED). Under the procedure of cooperative publications, it 
becomes in effect, a separate publication for each of the cooperating stations. 



Employment and program opportunities of cooperating institutions are offered to all people regardless of 
race, color, national origin, sex, age, or disability. 



Electronic Document Prepared by: Steven C. Hodges 



Publishing Institution: North Carolina State University 



Abstract 

The relative contribution of phosphorus (P) from agricultural nonpoint sources to 
surface water quality problems has increased in recent years as point sources of P have 
been reduced significantly. Phosphorus contributes to eutrophication, which restricts 
water use for fisheries, recreation, industry, and human consumption due to increased 
growth of undesirable algae and aquatic weeds, followed by oxygen shortages as the 
biomass decomposes. The increased attention on P has increased the demand for 
information on methods of analysis for soil, water, and residual materials for 
environmentally relevant forms of P. The purpose of this publication is to present these 
methods in a single document. Previously, the methods have appeared across a wide 
variety of documents or only in the scientific literature. It is not the intent of this 
publication to define a uniform set of recommended methods for agronomic soils tests, 
water, or residual materials. The methods presented here are intended solely to provide a 
set of uniform testing methods for environmental scientists working across an enormous 
range of soil and climatic conditions, with the hope that comparable methods may lead to 
improved communication and understanding of this complex issue. 



in 



FOREWARD 

As scientists focus on the fate of phosphorus applied to agricultural lands, it has become 
increasingly clear that a standard set of soil testing methods is needed to enable uniform 
comparison of results across county, state, regional, and even national boundaries. 

By contrast, soil testing developed with a high priority on meeting local needs. As a 
result, many local variations in extractants and laboratory procedures have been made to 
achieve timely analysis and improved correlation of soil test results with plant responses 
within well-defined regions. Over time, enormous amounts of information on individual 
soils, crops and extractants have been developed using these localized modifications and 
laboratory methods. Soil testing labs cannot easily change from one extractant to 
another. The cost of repeating these calibration experiments for many soils and crops is 
prohibitively expensive, and the changes would initially preclude users from comparing 
results across years. Even so, a set of standard reference methods can be useful for 
laboratories wishing to consider a new analysis for a particular element, and for 
comparing results across laboratories. In 1992, SERA-IEG-6 selected 15 reference 
procedures for soil testing laboratories in the southern region. Criteria for selection 
included the accuracy of the method in predicting crop responses, and general 
acceptability by workers in the soil testing field. 

This publication in no way attempts to define a uniform set of recommended methods for 
agronomic soil tests. The methods presented here are intended solely to provide a set of 
uniform testing methods for environmental scientists working across an enormous range 
of soil and climatic conditions, with the hope that comparable methods may lead to 
improved communication and understanding of this complex issue. 

For more information on agronomic soil testing methods, and the source of many of the 
procedures described here, the reader should refer to the recent bulletins compiled by the 
various regional committees working on nutrient analysis of soils, plants, water, and 
waste materials (SERA-IEG-6, NRC- 13 and NEC-67). 



IV 



CONTRIBUTORS 

Methods for Phosphorus Analysis for Soils, Sediments, Residuals, and Waters 



J. Thomas Sims 
Department of Plant & Soils 
531 South College Ave. 
University of Delaware 
Newark, DE 19717-1303 



D.A. Graetz 
Soil & Water Science 
P.O.Box 110510 
University of Florida 
Gainesville, FL 32611 



F.J. Coale 

Natural Resources Science 
H J. Patterson, Room 0225 
University of Maryland 
College Park, MD 20742-5821 

M.L. Self-Davis 

Crop, Soil & Environmental Science 
115 Plant Science Bldg. 
University of Arkansas 
Fayetteville, AR 72701-1201 

P. A. Moore 

Crop, Soil & Environmental Science 
115 Plant Science Bldg. 
University of Arkansas 
Fayetteville, AR 72701-1201 

B.C. Joern 

Department of Agronomy 

1150 Lilly Hall 

Purdue University 

West Lafayette, IN 47907-1 150 

W.J. Chardon 
P.O. Box 47 
6700 AC Wageningen 
The Netherlands 

O.F. Schoumans 

DLO Winand Staring Centre for 

Integrated Land, Soil & Water 

Research (SC-DLO) 

Dept. of Water and Environment 

P.O.Box 125 

6700 AC Wageningen 

The Netherlands 



V.D.Nair 

Soil Science Department 
106 Newell Hall 
University of Florida 
P.O.Box 110510 
Gainesville, FL 32611-0510 

A. Sharpley 

USDA-ARS 

Pasture System & Management 

Research 

Curtin Road 

University Park, PA 16802-3702 

M.R. Bender 

Department of Agronomy and Soils 

College of Agriculture 

202 Funchess Hall 

Auburn University, AL 36849-5412 



C.W.Wood 

Department of Agronomy and Soils 

College of Agriculture 

202 Funchess Hall 

Auburn University, AL 36849-5412 

H. Zhang 

Department of Plant Sciences 
Oklahoma State University 
Stillwater, OK 74078 

J.L. Kovar 

USDA/ARS National Soil Tilth 

Laboratory 

2150 Pammel Drive 

Ames, IA 50011-4420 



Thanh H. Dao 

USDA-ARS 

BARC - East Rm, 100B 

10300 Baltimore Ave., Bldg. 200 

Beltsville, MD 20705 

DA. Crouse 

Department of Soil Sciences 

Box 7619 

North Carolina State University 

Raleigh, NC 27695-7619 

S.C. Hodges 

Department of Soil Science 

Box 7629 

North Carolina State University 

Raleigh, NC 27695-7619 



A.C. Edwards 

Macaulay Land Use Research 

Institute 

Craigiebuckler 

Aberdeen 

AB15 8QH,UK 

D.H. Pote 

USDA/ARS 

5375 Hwy. 10 West 

Booneville, AR 72927 

T.C. Daniel 

Crop, Soil & Environmental Science 
115 Plant Science Bldg. 
University of Arkansas 
Fayetteville, AR 72701-1201 



C.R. Campbell 
NCDA Agronomic Division 
4300 Reedy Creed Road 
Raleigh, NC 27607-6465 

J.P. Zublena 

Cooperative Extension Service 

108 Ricks Hall 

North Carolina State University 

Raleigh, NC 27695-7602 

P.M. Haygarth 

Soil Science Group 

Institute of Grassland & 

Environmental Research 

North Wyke 

Okehampton 

Devon 

EX20 2SB, UK 



R.G. Myers 

Department of Agronomy 

2004 Throckmorton Plant Sciences 

Ctr. 

Kansas State University 

Manhattan, KS 66506-5501 

G.M. Pierzynski 

Department of Agronomy 

2004 Throckmorton Plant Sciences 

Ctr. 

Kansas State University 

Manhattan, KS 66506-5501 



VI 



Methods for P Analysis, G.M. Pierzynski (ed) 

TABLE OF CONTENTS 

Methods of Phosphorus Analysis for Soils, Sediments, Residuals, and 



Waters: Introduction. 



Gary M. Pierzynski, Kansas State University 
A.N. Sharpley, USDA-ARS, University Park, PA 



Soil Test Phosphorus: Principles and Overview 5 



J. Thomas Sims, University of Delaware 



Sample Collection, Handling, Preparation, and Storage 10 



Frank J. Coale, University of Maryland 



Soil Test Phosphorus: Bray and Kurtz P-l 13 



J. Thomas Sims, University of Delaware 



Soil Test Phosphorus: Mehlich 1 15 



J. Thomas Sims, University of Delaware 



Soil Test Phosphorus: Mehlich 3 17 



J. Thomas Sims, University of Delaware 



Soil Test Phosphorus: Olsen P 20 



J. Thomas Sims, University of Delaware 



A Phosphorus Sorption Index 22 



J. Thomas Sims, University of Delaware 



Determination of Water- and/or Dilute Salt-Extractable Phosphorus . 24 



M.L. Self-Davis, University of Arkansas 
PA. Moore, Jr., USDA-ARS, Fayetteville, AR 
B.C. Joern, Purdue University 
Phosphorus Extraction with Iron Oxide-Impregnated 



Filter Paper (P; test) 27 



W. J. Chardon, DLO Research Institute for Agrobiology and Soil Fertility, The 
Netherlands 
Determination of the Degree of Phosphate Saturation In Non- 



Calcareous Soils 31 



O.F. Schoumans, Winand Staring Centre for Integrated Land, Soil and Water 
Research, The Netherlands 



Phosphorus Sorption Isotherm Determination 35 



D.A. Graetz, University of Florida 
V.D. Nair, University of Florida 



Bioavailable Phosphorus in Soil 39 



Andrew Sharpley, USDA-ARS, University Park, PA 



Total Phosphorous in Soil 45 

M.R. Bender and C.W. Wood, Auburn University 



Phosphorus Fractionation 50 



Hailin Zhang, Oklahoma State University 
John L. Kovar, USDA/ARS, Ames, IA 



vn 



Methods for P Analysis, G.M. Pierzynski (ed) 



Phosphorus Fractionation in Flooded Soils and Sediments 60 



Philip Moore, USDA-ARS, Fayetteville, AR 
Frank Coale, University of Maryland 



Determination of Phosphorus Retention and Flux in Soil 65 



Thanh H. Dao, USDA-ARS 



Sampling Techniques for Nutrient Analysis of Animal Manures 71 



D.A. Crouse, S.C. Hodges, C.R. Campbell, J.P. Zublena, North Carolina State 
University 



Determining Water Soluble Phosphorus in Animal Manure 74 



M.L. Self-Davis, University of Arkansas 
PA. Moore, Jr., USDA-ARS, Fayetteville, AR 



Total Phosphorous in Residual Materials 77 



M.R. Bender and C.W. Wood, Auburn University 



Sample Collection, Handling, Preparation and Storage 84 



P.M. Haygarth, Institute of Grassland and Environmental Research, England 
A.C. Edwards, Macaulay Land Use Research Institute, Scotland 



Analyzing for Dissolved Reactive Phosphorus in Water Samples 91 



D.H. Pote, USDA-ARS, Booneville, AR 
T.C. Daniel, University of Arkansas 
Analyzing for Total Phosphorus and Total Dissolved Phosphorus in 



Water Samples 94 



D.H. Pote, USDA-ARS, Booneville, AR 
T.C. Daniel, University of Arkansas 
Using the Iron Oxide Method to Estimate Bioavailable Phosphorus in 



Runoff 98 



R.G. Myers, Kansas State University 
G.M. Pierzynski, Kansas State University 



Vlll 



Methods for P Analysis, G.M. Pierzynski (ed) 

Methods of Phosphorus Analysis for Soils, Sediments, 
Residuals, and Waters: Introduction 

Gary M. Pierzynski, Kansas State University 

A.N. Sharpley, USDA-ARS, University Park, PA 

Point sources of water pollution have been reduced significantly since the late 1960s 
due to their relative ease of identification, legislation, and advances in pollution control 
technology. Consequently, the relative contribution of agricultural nonpoint sources to 
remaining water quality problems has increased. Of the water quality issues that remain, 
a recent EPA survey has identified eutrophication as the single largest problem in surface 
water quality. 

Eutrophication restricts water use for fisheries, recreation, industry, and drinking, due 
to increased growth of undesirable algae and aquatic weeds, followed by oxygen 
shortages as the biomass decomposes. Also, many drinking water supplies throughout 
the world undergo periodic massive surface blooms of cyanobacteria. These blooms 
contribute to a wide range of water-related problems, including summer fish kills, 
unpalatability of drinking water, and formation of trihalomethane, a known carcinogen, 
during water chlorination. Recent outbreaks of the dinoflagellate Pfiesteria piscicida in 
the eastern U.S. have also been linked to excess nutrients in affected waters. 
Neurological damage in people exposed to the highly toxic volatile chemical produced by 
this dinoflagellate has dramatically increased public awareness of eutrophication and the 
need for solutions. In most cases, phosphorus (P) accelerates the eutrophication of fresh 
waters. Consequently, controlling algal blooms and eutrophication mainly requires 
reducing P inputs to surface waters. 

The purpose of this manual is to present methods for analysis of soil, water, and 
residual materials for environmentally relevant P forms in a single document. Previously, 
these methods appeared separately in methods publications for soil or water or have only 
appeared in the scientific literature. Commercial and research laboratories today must 
deal with the analysis of a wider range of sample types for more diverse agronomic and 
environmental uses. This has caused confusion over selection of the most appropriate 
method for a specific need and can lead to inappropriate recommendations for P 
management. Thus, there is an urgent need for a publication containing all of the 
currently available procedures for P analysis. 

The mainstay of P analysis for all solution types has been use of colorimetric 
procedures, most notably from Murphy and Riley (1962). Colorimetric procedures are 
sensitive, reproducible, and lend themselves to automated analysis. In addition, the 
methods can accommodate water samples, digest solutions, and extracts. The basic 
Murphy and Riley procedure is presented in Sharpley (2000) in this bulletin. Variations 
in the procedure are incorporated into other sections, despite the appearance of 
redundancy. Modifications to the procedures are often method- specific. 

Inductively coupled plasma (ICP) spectrophotometry can also be used for P 
determination. The use of ICP has increased as the use of multi-element soil extractants 
becomes more popular. Results from colorimetric analyses are not always directly 
comparable to those from ICP because ICP estimates the total amount of P in solution, 



Methods for P Analysis, G.M. Pierzynski (ed) 

while the colorimetric procedures measure P that can react with the color developing 
reagent. 

Nomenclature for forms of P in soil, water, or residual materials varies in the 
literature, particularly for operationally-defined forms of P in water samples. Table 1 
presents an abbreviated description of forms of P in runoff or drainage water that have 
been used in the literature and that we propose as a standardized terminology. 
Phosphorus forms in soils are also difficult to standardize with any reasonable consensus, 
due to the number of different disciplines involved (e.g., soil scientists, agronomists, 
limnologists, hydrologists). Thus, beyond using total soil P, we strongly encourage the 
use of specific chemical terminology (e.g., water extractable, CaCl2 extractable, 0.1 M 
NaOH extractable, Mehlich extractable P, etc.), which has been clearly defined. Any 
other terminology, which may be used in conclusions and interpretations (e.g., 
desorbable, available, bioavailable, sorbed P etc.), must also be clearly defined. 

Traditionally, extractable P has been used by soil testing laboratories to describe the 
amount of P in soil available for crop uptake and to determine the probability of crop 
response to added P, and thereby fertilizer P requirements. Bioavailable P is often used 
to describe P in soil or sediment that is available for uptake by algae or macrophytes in 
surface waters. Occasionally, bioavailable P is used to describe the availability of soil P 
to plants. There are also a large number of soil P extraction methods that have been 
designed to account for various soil types and mechanisms controlling the chemistry of 
soil P. For example, numerous soil extractants are available for acid soils, where Al and 
Fe dominate P chemistry, and basic or calcareous soils, where Ca dominates soil P 
reactions. 

Clearly, there is a potential for confusion by the uninitiated. Hence it is essential to 
accurately define how P was measured in soil or water samples to avoid potential 
misinterpretations or inappropriate recommendations. This publication documents in 
detail the analytical methods available, their recommended uses, and some information 
on interpretation. 



References: 

Murphy, J., and J. P. Riley. 1962. A modified single solution method for determination of 
phosphate in natural waters. Anal. Chim. Acta. 27:31-36. 

Sharpley, A.N. 2000. Bioavailable phosphorus in soil. In G.M. Pierzynski (ed.), Methods 
for Phosphorus Analysis for Soils, Sediments, Residuals, and Waters. Southern 
Cooperative Series Bulletin No. 396, p. 39-45. 



Methods for P Analysis, G.M. Pierzynski (ed) 



Table 1. Proposed standardization of terminology for forms of P in runoff and drainage 
water. 



Phosphorus Form 



Abbreviation Example Methodology 



r 



Total Phosphorus 

Total amount in dissolved 
and particulate phases 



Total Dissolved Phosphorus 
Dissolved inorganic (ortho 
P) and organic P 

Dissolved Orthophosphate 

Immediately algal available 

Bioavailable Phosphorus 

Dissolved ortho P and a 
portion of particulate P that 
is algal available 



Molybdate Reactive Phosphorus 

Dissolved ortho P and acid 
extractable particulate P 
(possibly algal available) 

Particulate Phosphorus 

Inorganic and organic P 
associated with or bound to 
eroded sediment 

Dissolved Organic Phosphorus:]: 

Includes polyphosphates and 
hydrolyzable phosphates 



TP 


Digestion of unfiltered water 




sample 




-Kjeldahl procedure 




-Acid ammonium persulfate 




-Perchloric acid 


TDP 


Acid persulfate digestion of 




unfiltered sample 


DP 


Murphy and Riley on filtered 




sample 


BAP 


Extraction of unfiltered samp 



MRP 



PP 



DOP 



with 
-NaOH 

-NaCl 

Anion exchange resin 

-Ammonium fluoride 

-Iron-oxide filter paper strips 

Murphy and Riley colorimetric 
analysis of an unfiltered sample 



By difference = [TP - TDP] 



By difference = [TDP - DP] 



f Not an inclusive list of appropriate methods that can be used. Filtered samples 

are defined as that passing through a 0.45um filter. 
$ If dissolved organic P constitutes more than 25% of TDP, then measuring 

polyphosphates and hydrolyzable phosphates may be necessary. 



Methods for P Analysis, G.M. Pierzynski (ed) 



Soils and Sediments 



Methods for P Analysis, G.M. Pierzynski (ed) 

Soil Test Phosphorus: Principles and Overview 
J. Thomas Sims, University of Delaware 



Principles of Soil Testing for Phosphorus: 

Soil testing for phosphorus (P) has been formally conducted in the United States since 
the late 1940s and is now a well-established agronomic practice. The fundamental goal 
of soil P testing has always been to identify the "optimum" soil test P concentration 
required for plant growth. The need for additional fertilization or manuring, and the 
economic return on an investment in fertilizer P, could then be predicted. Sims et al. 
(1998) stated that other objectives of soil P testing have been to: (i) "index" the P 
supplying capacity of soils, thus estimating the time before fertilization would again be 
required; (ii) group soils, in terms of the likelihood of an economic response to P, based 
on their physical and chemical properties; and, (iii) most recently, to identify when soils 
are sufficiently excessive in P to contribute to nonpoint source pollution of surface 
waters. Bray (1948) proposed that an acceptable agronomic soil P test should have the 
following characteristics: 

• The soil test should extract all or a proportionate amount of the plant-available 
P from soils with differing chemical and mineralogical properties. 

• The soil test should be accurate and rapid. 

• The P extracted by the soil test should be well correlated with plant P 
concentration, plant growth, and the response of the plant to added P in 
fertilizers or manures. 

• The soil test should accurately detect differences in soil P concentrations 
caused by previous fertilization or manuring. 

The major steps involved in a soil P testing program are outlined in Table 1 (from 
Sims et al., 1998). From an agronomic perspective, if these steps are followed, soil P 
management will be successful and economically beneficial. However, if the goal of soil 
P testing is to assess the potential environmental impact of soil P, a thorough re-analysis 
of each step in the soil testing process, from sample collection to interpretation of results 
should be conducted. Several recent reviews address the principles and practices 
involved in environmental soil testing for P (Sibbesen and Sharpley, 1997; Sims, 1993; 
Sims, 1997; Sims, 1998; Sims et al., 2000). 

The purpose of the following sections is to provide an overview of the four soil test P 
methods most commonly used in the United States and Canada today (Bray and Kurtz P- 
1, Mehlich 1, Mehlich 3, and Olsen P). Detailed descriptions of the laboratory methods 
and analytical procedures used to determine P by these methods are provided in other 
references (Carter, 1993; Frank, et al, 1998; Kuo, 1996; SERA-IEG-6, 1992; Sims and 
Wolf, 1995; SPAC, 1992). Finally, Table 2 lists other soil test P methods now used 
domestically and in other countries, and provides references for each method. 



Methods for P Analysis, G.M. Pierzynski (ed) 



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Methods for P Analysis, G.M. Pierzynski (ed) 

References: 

Bray, R.H. 1948. Correlation of soil tests with crop response to fertilizers and with 

fertilizer requirement, p. 53-86. In H.B. Kitchen (ed.) Diagnostic techniques for soils 

and crops. Am. Potash Inst., Washington, D. C. 
Carter, M.R. (ed.) 1993. Soil Sampling and Methods of Analysis. Can. Soc. Soil Sci., 

Ottawa, Ontario, Canada. 
Chardon, W.J., R.G. Menon, and S.H. Chien. 1996. Iron oxide impregnated filter paper 

(Pi test): A review of its development and methodological research. Nutr. Cycl. 

Agroecosystems 46:41-51. 
Egner, H., H. Riehm, and W.R. Domingo. 1960. Untersuchungen uber die chemishe 

bodenanalyse als grundlage fur die beurteilung des nahrstoffzustandes der boden. H 

Chemische extraktions-methoden zu phosphor - and kalimbestimmung kungl. 

Lantbrukshoegsk. Ann. 26:204-209. 
Houba, V.J.G., J.J. van der Lee, and I. Novozamski. 1997. Soil analysis procedures. 

Dep. of Soil Sci. Plant Nutr., Landbouwuniversiteit, Wageningen Agric. Univ., 

Wageningen, the Netherlands. 
Frank, K., D. Beegle, and J. Denning. 1998. Phosphorus, p. 21-30. In J.R. Brown (ed.) 

Recommended Chemical Soil Test Procedures for the North Central Region. North 

Central Reg. Res. Publ. No. 221 (revised). 
Kuo, S. 1996. Phosphorus, p. 869-919. In D. L. Sparks, (ed.) Methods of Soil Analysis: 

Part 3- Chemical Methods. SSSA, Madison, WI. 
Lunt, H. A., C. L.W. Swanson, and H.G.M. Jacobson. 1950. The Morgan Soil Testing 

System. Bull. No. 541, Conn. Agr. Exp. Stn., New Haven, CT. 
Mcintosh, J. L. 1969. Bray and Morgan soil test extractants modified for testing acid 

soils from different parent materials. Agron. J. 61:259-265. 
Menon, R.G., S.H. Chien, and W.J. Chardon. 1997. Iron oxide impregnated filter paper 

(Pi test): II. A review of its application. Nutr. Cycl. Agroecosystems 47:7-18. 
Morgan, M.F. 1941. Chemical soil diagnosis by the universal soil testing system. Conn. 

Agric. Exp. Stn. Bull. No. 450. 
Meyers, R.G., G.M. Pierzynski, and S.J. Thien. 1995. Improving the iron oxide sink 

method for extracting soil phosphorus. Soil Sci. Soc. Am. J. 59:853-857. 
SERA-IEG-6 (Southern Extension Research Activity - Information Exchange Group) 

1992. Donohue, S.J. (ed.) Reference Soil and Media Diagnostic procedure for the 

southern region of the United States. So. Coop. Series Bulletin 374. Va. Agric. Exp. 

Station, Blacksburg, VA. 
Sibbesen, E., and A.N. Sharpley. 1997. Setting and justifying upper critical limits for 

phosphorus in soils. In H. Tunney et al., (ed.) Phosphorus Loss from Soil to Water. 

CAB International, London. 
Sims, J.T. 1993. Environmental soil testing for phosphorus. J. Prod. Agric. 6:501-507. 
Sims, J.T. 1997. Phosphorus soil testing: Innovations for water quality protection, p. 47- 

63. Proc. 5th Intl. Symp. Soil and Plant Analysis. Minneapolis, MN. 
Sims, J.T. (ed.) 1998. Soil testing for phosphorus: Environmental uses and implications. 

So. Coop. Series Bull. No. 389. Univ. Delaware, Newark, DE. 
Sims, J.T., and A.M. Wolf, (ed.) 1995. Recommended soil testing procedures for the 

Northeastern United States. (2nd ed.). Bull. No. 493. Univ. Delaware, Newark, DE. 



Methods for P Analysis, G.M. Pierzynski (ed) 

Sims, J.T., A.C. Edwards, O.F. Schoumans, and R.R. Simard. 2000. Integrating soil 
phosphorus testing into environmentally based agricultural management practices. J. 
Environ. Qual. In Press. 

Sims, J.T., S.C. Hodges, and J. Davis. 1998. Soil testing for phosphorus: Current status 
and uses in nutrient management programs, p. 13-20. In Sims, J.T. (ed.) 1998. Soil 
testing for phosphorus: Environmental uses and implications. So. Coop. Series Bull. 



No. 389. Univ. Delaware, Newark, DE. ( ittp://www.soil.ncsu.edu/seral7/ ) 



Sissingh, H.A. 1971. Analytical technique of the Pw method used for the assessment of 

the phosphate status of arable soils in the Netherlands. Plant Soil 34:438-446. 
SPAC (Soil and Plant Analysis Council). 1992. Handbook on reference methods for soil 

analysis. Georgia Univ. Stn., Athens, GA. 
Soltanpour, P.N. and A.P. Schwab. 1977. A new soil test for simultaneous determination 

of macro and micro-nutrients in alkaline soils. Commun. Soil Sci. Plant Anal. 8:195- 

207. 



Methods for P Analysis, G.M. Pierzynski (ed) 

Sample Collection, Handling, Preparation, and Storage 
Frank J. Coale, University of Maryland 

Sample collection: 

The collection of a representative and reliable soil sample for phosphorus (P) analysis 
requires predetermination of sampling depth, position relative to nutrient application 
patterns, and sampling intensity within the field. The appropriate soil sampling depth is 
dependent upon the planned interpretation of the analytical data. If investigation of P 
distribution or concentration with depth is a specified research objective, three factors 
must be considered when determining the appropriate sampling depth: 1) influence of 
changes in soil morphology with depth (i.e., horizonation); 2) influence of surface soil 
management (e.g., tillage); and 3) necessity to maintain sample collection depth 
uniformity across numerous sites. 

Sample collection depth based on observed morphological horizon depths is quite 
useful when attempting to associate soil P measurements with soil physical properties. 
This technique may generate very reliable data for a particular, well-defined location, but 
this laborious task is not very practical when a research project focuses on more than a 
few soils or when the data will be subjected to broader, perhaps watershed-scale, 
interpretation. 

Depth of tillage will dramatically impact soil P distribution with depth. Tillage depth 
is seldom constant across a given field. Sampling depths should include soil collected 
from a depth confidently within the tillage zone and excluding soil from below the tillage 
zone. A second transitional depth should be collected that is expected to be variably 
affected by tillage and includes the lower tillage boundary. Deeper sampling depths 
should not be directly impacted by physical tillage activity. 

Relating soil physical and chemical properties to the potential for P transport with 
surface runoff water requires a different approach to soil sample collection. Sharpley 
(1985) studied five soils of varying physical and chemical properties and found that 
effective depth of interaction between surface soil and runoff ranged from 2 to 40 mm. 
The effective depth of interaction varied by soil type, surface slope, rainfall intensity, and 
crop residue. For most agricultural soils, samples collected to a depth of 20 mm would 
accurately define the effective depth of runoff interaction generated by moderate to high 
rainfall intensity (< 50 mm/h). For medium to coarse textured soils on steeper slopes 
(>12 %) that are subjected to high intensity rainfall (> 100 mm /h), soils should be 
sampled to a depth of 40 mm in order to more accurately relate the potential for P 
transport with surface runoff to soil physical and chemical properties. 

Recommended soil sampling intensity is usually between 10 and 30 subsamples per 
composite sample (Whitney et al., 1985; Kitchen et al., 1990; Coale, 1997). A single 
composite sample may represent a single research plot or an entire production field, but 
generally not more than 10 ha. 

Discrete nutrient application patterns in a field can increase the complexity of 
appropriate soil sample collection procedures. In a review of positional P availability 
resulting from band application of fertilizer P, Sharpley and Halvorson (1994) stated that 
collection of 15 random samples (Ward and Leikam, 1986; Shapiro, 1988) to 30 random 
samples (Hooker, 1976) were adequate to reflect crop P availability in conventionally 



10 



Methods for P Analysis, G.M. Pierzynski (ed) 

tilled fields where previous P fertilizer bands exist. For no-till or minimum-till soils 
containing residual P fertilizer bands in which the location of the P bands is known, 
sampling to include one "in-the-band" soil sample for every 20 "between-the-band" 
samples for 76 cm band spacing, and one "in-the-band" sample for every 8 "between-the- 
band" samples for 30 cm band spacing, will accurately reflect the mean soil P status of 
the field (Kitchen et al., 1990). Twenty to 30 subsamples per composite are adequate. 
When the location of the P bands is not known, collection of 20 to 30 subsamples per 
composite is also adequate but paired subsamples should be collected where the location 
of the first subsample of the pair is completely random and the second subsample of the 
pair is located 50% of the band-spacing distance from the first, perpendicular to the band 
direction (Kitchen et al, 1990). 



Sample Handling, and Preparation and Storage: 

Air-drying should be satisfactory for investigations into relative changes in soil P 
concentrations in response to imposed treatments or for routine comparative P analyses. 
Soil samples should be air-dried (25 to 30°C) and crushed to pass a 2 mm sieve. Air- 
dried and crushed soil samples are stable at room temperature. Air-drying may not be 
suitable for determination of the absolute quantity of the various P fractions in soils. Air- 
drying may artificially elevate the quantity of soluble reactive P above in situ conditions. 
Bartlett and James (1980) studied P solubility in the surface soil of a loamy fine sand and 
found water-soluble P concentrations to be five times higher in air-dried samples (-30 
mg P/ L) than in samples stored at field moisture (~5 mg P/ L). The effect of air-drying 
was only partially reversed by rewetting and incubating the air-dried soil for one month 
(-20 mg P /L). Water-soluble P in re wetted soil samples that had previously been air- 
dried was shown to decrease during three months of storage at 20° C (Bartlett and James, 
1980). For quantitative characterization studies, soil and sediment samples should be 
stored at field moisture content under refrigeration, between and 4°C. Soil and 
sediment samples should not be stored frozen (<0°C), because the water-soluble 
proportion of total P increases after freezing (Mack and Barber, 1960). Mixing moist soil 
samples to achieve homogeneity is difficult, and careful attention should be paid to 
ensure thorough mixing prior to subsampling. Moist soils are also difficult to sieve, but 
large particles (> 2mm) should be removed from the sample prior to analysis. 



References: 

Bartlett, R. and B. James. 1980. Studying dried, stored soil samples - some pitfalls. Soil 

Sci. Soc. Am. J. 44:721-724. 
Coale, F. J. (ed.) 1997. Chesapeake Bay region nutrient management training manual. 

USEPA Chesapeake Bay Program, Annapolis, MD. 
Hooker, M.L. 1976. Soil sampling intensities required to estimate available N and P in 

five Nebraska soil types. MS Thesis, Univ. Nebraska, Lincoln, NE. 
Kitchen, N.R., J.L. Havlin, and D.G. Westfall. 1990 Soil sampling under no-till banded 

phosphorus. Soil Sci. Soc. Am. J. 54:1661-1995. 



11 



Methods for P Analysis, G.M. Pierzynski (ed) 

Mack, A.R. and S.A. Barber. 1960. Influence of temperature and moisture on soil 

phosphorus. I. Effect on soil phosphorus fractions. Soil Sci. Soc. Am. Proc. 24:381- 

385. 
Shapiro, C.A. 1988. Soil sampling fields with a history of fertilizer bands. Soil Sci. 

News, Nebraska Coop. Ext. Serv., Vol. 10, No. 5. 
Sharpley, A.N. 1985. Depth of surface soil-runoff interaction as affected by rainfall, soil 

slope, and management. Soil Sci. Soc. Am. J. 49:1010-1015. 
Sharpley, A.N. and A.D. Halvorson. 1994. The management of soil phosphorus 

availability and its impact on surface water quality, p. 7-90. In R. Lai and B. A. 

Stewart (ed.) Soil processes and water quality. Advances in Soil Science. Lewis 

Publishers, Boca Raton, FL. 
Ward, R. and D.F. Leikman. 1986. Soil sampling techniques for reduced tillage and band 

fertilizer application. In Proc. Great Plains Soil Fertility Workshop. March 4-5, 

1986. Denver, CO. 
Whitney, DA., J.T. Cope, and L.F. Welch. 1985. Prescribing soil and crop nutrient 

needs, p. 25-52. In O. P. Engelstad (ed.) Fertilizer technology and use. 3 rd ed. SSSA, 

Madison, WI. 



12 



Methods for P Analysis, G.M. Pierzynski (ed) 

Soil Test Phosphorus: Bray and Kurtz P-l 
J. Thomas Sims, University of Delaware 

Introduction: 

The Bray and Kurtz P-l soil test phosphorus (P) method was developed by Roger H. 
Bray and Touby Kurtz of the Illinois Agricultural Experiment Station in 1945 and is now 
widely used in the Midwestern and North Central United States (Bray and Kurtz, 1945; 
Frank et al., 1998). Phosphorus extracted by the Bray and Kurtz P-l method has been 
shown to be well-correlated with crop yield response on most acid and neutral soils in 
these regions. For acid soils, the fluoride in the Bray and Kurtz extractant enhances P 
release from aluminum phosphates by decreasing Al activity in solution through the 
formation of various Al-F complexes. Fluoride is also effective at suppressing the re- 
adsorption of solubilized P by soil colloids. The acidic nature of the extractant (pH 2.6) 
also contributes to dissolution of available P from Al, Ca, and Fe-bound forms in most 
soils. The Bray soil test is not suitable for: 

• clay soils with a moderately high degree of base saturation, 

• silty clay loam or finer-textured soils that are calcareous or have a high pH 
value (pH > 6.8) or have a high degree of base saturation, 

• soils with a calcium carbonate equivalent > 7% of the base saturation, or 

• soils with large amounts of lime (> 2% CaCCh). 

In soils such as these, the acidity of the extracting solution can be neutralized unless 
the ratio of extractant: soil is increased considerably. Additionally, CaF 2 , formed from the 
reaction of soluble Ca + in the soil with F- added in the extractant, can react with and 
immobilize soil P. Both types of reactions reduce the efficiency of P extraction and result 
in low soil test P values. Finally, the Bray and Kurtz extractant can dissolve P from rock 
phosphates, therefore it should not be used in soils recently amended with these 
materials, as it will overestimate available P. A Bray and Kurtz P-l value of 25 to 30 mg 
P/kg soil is often considered optimum for plant growth, although Holford (1980) reported 
lower critical values for highly buffered soils. 

Equipment: 

1. No. 10 (2 mm opening) sieve 

2. Standard 1 g and 2 g stainless steel soil scoops 

3. Automatic extractant dispenser, 25 mL capacity 

4. Extraction vessels, such as 50 mL Erlenmeyer flasks, and filter funnels (9 and 1 1 
cm) and racks 

5. Rotating or reciprocating shaker with a capability of 200 excursions per minute 
(epm) 

6. Whatman No. 42 or No. 2 (or equivalent) filter paper, 9 to 1 1 cm. (Acid resistant 
filter paper may be needed if using an automated method for determining P 
concentration by intensity of color. Bits of filter paper may cause an obstruction in 
the injection valves.) 



13 



Methods for P Analysis, G.M. Pierzynski (ed) 

Reagents: 

1. Bray and Kurtz P-l Extracting Solution (0.025 M HC1 in 0.03 M NH4F): Dissolve 
1 1. 1 1 g of reagent-grade ammonium fluoride (NH4F) in about 9 L of distilled water. 
Add 250 mL of previously standardized 1M HC1 and make to 10 L volume with 
distilled water. Mix thoroughly. The pH of the resulting solution should be pH 2.6 + 
0.05. The adjustments to pH are made using HC1 or ammonium hydroxide 
(NH40H). Store in polyethylene carboys until use. 

Procedure: 

1. Scoop or weigh 2 g of soil into a 50 mL Erlenmeyer flask, tapping the scoop on the 
funnel or flask to remove all of the soil from the scoop. 

2. Add 20 mL of extracting solution to each flask and shake at 200 or more epm for five 
minutes at a room temperature at 24 to 27°C 

3. If it is necessary to obtain a colorless filtrate, add 1 cm 3 (-200 mg) of charcoal 
(DARCO G60, J. T. Baker, Phillipburg, NJ) to each flask. 

4. Filter extracts through Whatman No. 42 filter paper or through a similar grade of 
paper. Refilter if extracts are not clear. 

5. Analyze for P by colorimetry or inductively coupled plasma emission spectroscopy 
using a blank and standards prepared in the Bray P-l extracting solution. 

Calculations: 

Bray and Kurtz P-l Extractable phosphorus is calculated as 

t, 1T . , i tw ™ -,x C p x [0.020 L extract] 
Bray and Kurtz P - 1 Extractable P (mg P/kg soil ) = — 

0.002 kg soil 
where 

Cp = Concentration of P in Bray and Kurtz P-l extract, in mg/L . 



References: 

Bray R.H., and L.T. Kurtz. 1945. Determination of total, organic and available forms of 

phosphorus in soils. Soil Sci. 59:39-45. 
Frank, K.D. Beegle, and J. Denning. 1998. Phosphorus, p. 21-30. In J. R. Brown (ed.) 

Recommended Chemical Soil Test Procedures for the North Central Region. North 

Central Reg. Res. Publ. No. 221 (revised). 
Holford, I.C.R. 1980. Greenhouse evaluation of four phosphorus soil tests in relation to 

phosphate buffering and labile phosphate in soils. Soil Sci. Soc. Am. J. 44:555-559. 



14 



Methods for P Analysis, G.M. Pierzynski (ed) 

Soil Test Phosphorus: Mehlich 1 
J. Thomas Sims, University of Delaware 



Introduction: 

The Mehlich 1 soil test for phosphorus (P), also known as the dilute double acid or 
North Carolina extractant, was developed in the early 1950s by Mehlich and his co- 
workers (Mehlich, 1953; Nelson et al. 1953). In the United States the Mehlich 1 
procedure is primarily used in the southeastern and mid- Atlantic states as a multi-element 
extractant for P, K, Ca, Mg, Cu, Fe, Mn, and Zn. The Mehlich 1 extracts P from 
aluminum, iron, and calcium phosphates and is best suited to acid soils (pH < 6.5) with 
low cation exchange capacities (< 10 cmol/kg) and organic matter contents (< 5%). Kuo 
(1996) reported that the Mehlich 1 soil test was unreliable for calcareous or alkaline soils 
because it extracts large amounts of nonlabile P in soils with pH > 6.5, soils that have 
been recently amended with rock phosphate, and soils with high cation exchange capacity 
(CEC) or high base saturation. In soils such as these the acidity of the Mehlich 1 solution 
is neutralized, reducing the capability of the dilute acid to extract P. Similar reductions in 
P extraction efficiency have been attributed to clay and hydrous aluminum and iron 
oxides (Nelson et al., 1953; Lins & Cox, 1989). 

A Mehlich 1 P value of 20 to 25 mg P/kg soil for the Mehlich- 1 test is generally 
considered to be optimum for plant growth, although this may vary slightly between soil 
types and cropping systems. For instance, Kamprath and Watson (1980) stated a 
Mehlich- IP of 20 to 25 mg P/kg soil is adequate for plants grown in sandy soils but only 
10 mg P/kg soil is required for fine-textured soils, a point supported by the work of Lins 
and Cox (1989). 

Equipment: 

1. No. 10 (2 mm opening) sieve 

2. Automatic extractant dispenser, 25 mL capacity (If preferred, pipettes are 
acceptable.) 

3. Standard 5 cm 3 and 1 cm 3 stainless steel soil scoops 

4. Extraction vessels, such as 50 mL Erlenmeyer flasks, and filter funnels (9 and 1 1 
cm) and racks 

5. Reciprocating or rotary shaker, capable of at least 180 epm (excursions per 
minute) 

6. Whatman No. 42 or No. 2 (or equivalent) filter paper, 9 to 1 1 cm. (Acid resistant 
filter paper may be needed if using an automated method for determining P 
concentration by intensity of color. Bits of filter paper may cause an obstruction in 
the injection valves.) 

Reagents: 

1. Mehlich 1 Extracting Solution (0.0125 M H2S04 + 0.05 M HC1). Also referred 
to as dilute double acid or the North Carolina Extractant. Using a graduated 
cylinder, add 167 mL of concentrated HC1 (12M) and 28 mL of concentrated 
H2S04 (18M) to -35 L of deionized water in a large polypropylene carboy. 



15 



Methods for P Analysis, G.M. Pierzynski (ed) 

Make to a final volume of 40 L by adding deionized water. Mix well by bubbling 
air through the solution for 3 hours. 

Procedure: 

7. Weigh 5.0 g (or scoop 4 cm 3 ) of sieved (< 2 mm), air-dried soil into a 50 mL 
extraction flask. 

8. If it is necessary to obtain a colorless filtrate, add 1 cm3 (-200 mg) of charcoal 
(DARCO G60, J. T. Baker, Phillipburg, NJ) to each flask. 

9. Add 20 mL of the Mehlich 1 extracting solution and shake for five minutes on a 
reciprocating shaker set at a minimum of 180 epm at a room temperature at 24 to 
27°C. 

10. Filter through a medium-porosity filter paper (Whatman No. 2 or equivalent). 

11. Analyze for P by colorimetry or inductively coupled plasma emission 
spectroscopy using a blank and standards prepared in the Mehlich 1 extracting 
solution. 

Calculations: 

Mehlich 1 Extractable P (mg P/kg soil) = 

[Concentration of P in Mehlich 1 extract, mg/L ] x [ 0.020 L extract -f 0.005 kg 
soil] 



References: 

Kamprath, E.J. and M.E. Watson. 1980. Conventional soil and tissue tests for assessing 

the phosphorus status of soils, p. 433-469. In F. E. Khasawneh et al. (ed.) The role 

of phosphorus in agriculture. ASA, CSSA, and SSSA, Madison, WI. 
Kuo, S. 1996. Phosphorus, p. 869-919. In D. L. Sparks, (ed.) Methods of Soil Analysis: 

Part 3- Chemical Methods. SSSA, Madison, WI. 
Lins, I.D.G. and F.R. Cox. 1989. Effects of extractant and selected soil properties on 

predicting the optimum phosphorus fertilizer rate for growing soybeans under field 

conditions. Commun. Soil Sci. Plant Anal. 20:310-333. 
Mehlich, A. 1953. Determination of P, Ca, Mg, K, Na, and NFL. North Carolina Soil 

Test Division (Mimeo). Raleigh, NC. 
Nelson, W. L., A. Mehlich, and E. Winters. 1953. The development, evaluation, and use 

of soil tests for phosphorus availability. Agronomy 4:153-158. 



16 



Methods for P Analysis, G.M. Pierzynski (ed) 

Soil Test Phosphorus: Mehlich 3 
J. Thomas Sims, University of Delaware 



Introduction: 

The Mehlich 3 soil test was developed by Mehlich in 1984 as an improved multi- 
element extractant for P, K, Ca, Mn, Cu, Fe, Mn, and Zn (Mehlich, 1984). Today, the 
Mehlich 3 test is used throughout the United States and Canada because it is well suited 
to a wide range of soils, both acidic and basic in reaction. The Mehlich 3 extractant was 
selected by workers in the southern region as the standard reference procedure for soil 
test P determination (Tucker, 1992). The Mehlich 3 is similar in principle to the Bray 
and Kurtz P-l test because it is an acidic solution that contains ammonium fluoride. 
Acetic acid in the extractant also contributes to the release of available P in most soils. It 
is more effective than the Mehlich 1 soil test at predicting crop response to P on neutral 
and alkaline soils because the acidity of the extractant is neutralized less by soil 
carbonates (Tran and Simard, 1993). Several studies showed that the Mehlich 3 soil test 
is highly correlated with P extracted from soils by the Bray and Kurtz P-l, Mehlich 1, 
and Olsen P methods (Sims, 1989; Tran et al, 1990; Wolf and Baker, 1985). 

A Mehlich 3 value of 45-50 mg P/kg soil is generally considered to be optimum for 
plant growth and crop yields, higher than the critical values used for other standard soil P 
tests such as the Bray and Kurtz P-l, Mehlich 1, and Olsen P. 

Equipment: 

1. No. 10 (2 mm opening) sieve 

2. Standard 1 cm 3 , 2 cm 3 (or 2.5 cm 3 ) stainless steel soil scoops 

3. Automatic extractant dispenser, 25 mL capacity 

4. Extraction vessels, such as 50 mL Erlenmeyer flasks, and filter funnels (9 and 1 1 
cm) and racks 

5. Rotating or reciprocating shaker with a capability of 200 excursions per minute 
(epm) 

6. Whatman No. 42 or No. 2 (or equivalent) filter paper, 9 to 1 1 cm. (Acid resistant 
filter paper may be needed if using an automated method for determining P 
concentration by intensity of color. Bits of filter paper may cause an obstruction in 
the injection valves.) 



Reagents: 

1. Mehlich 3 Extracting Solution: (0.2 M CH3COOH, 0.25 M NH4N03, 0.015 M 
NH4F, 0.013 M HN03, 0.001 M EDTA [(HOOCCH2)2NCH2CH2N 
(CH2COOH)2]. Prepare as follows: 

Ammonium fluoride (NH4F) and EDTA stock solution (3.75 M NH4F:0.25 M EDTA) 

2. Add 1,200 mL of distilled water to a 2 L volumetric flask. 

3. Add 277.8 g of NH4F and mix well. 

4. Add 146. 1 g EDTA to the solution. 



17 



Methods for P Analysis, G.M. Pierzynski (ed) 

5. Make solution to 2 L, mix well and store in plastic (stock solution for 10,000 
samples). 

Mehlich 3 extractant preparation 

6. Add 8 L of distilled water to a 10 L carboy. 

7. Dissolve 200 g of ammonium nitrate (NH4N03) in the distilled water. 

8. Add 40 mL NH4F-EDTA stock solution and mix well. 

9. Add 1 15 mL glacial acetic acid (99.5%, 17.4 M). 

10. Add 8.2 mL of concentrated nitric acid (HN03, 68 to 70 %, 15.5 M). 

11. Add distilled water to 10 L final volume and mix well (enough extractant for 400 
samples), final pH should be 2.5 ± 0.1. 

Procedure: 

1. Scoop or weigh 2.0 g of soil into a 50 mL Erlenmeyer flask, tapping the scoop on 
the funnel or flask to remove all of the soil from the scoop. Where disturbed bulk 
density of soil varies significantly from 1.0 g cm 3 , record both weight and volume 
of samples. (Standard 2.5 cm 3 scoops may also be used, but a 1:10 soil: extractant 
volumetric ratio should be maintained) 

2. Add 20 mL of extracting solution to each flask and shake at 200 or more epm for 
five minutes at a room temperature at 24 to 27°C. 

3. If it is necessary to obtain a colorless filtrate, add 1 cm 3 (-200 mg) of charcoal 
(DARCO G60, J. T. Baker, Phillipburg, NJ) to each flask. 

4. Filter extracts through Whatman No. 42 filter paper or through a similar grade of 
paper. Refilter if extracts are not clear. 

5. Analyze for P by colorimetry or inductively coupled plasma emission 
spectroscopy using a blank and standards prepared in the Mehlich 3 extracting 
solution. 

Calculations: 

Mehlich 3 Extractable P (mg P/kg) = 

[Concentration of P in Mehlich 3 extract, mg P/L] x [ 0.020 L extract 4- 0.002 kg soil] 



References: 

Mehlich, A. 1984. Mehlich 3 soil test extractant: A modification of the Mehlich 2 

extractant. Commun. Soil Sci. Plant Anal. 15:1409-1416. 
Tucker, M.R., 1992. Determination of phosphorus by Mehlich 3 extractant. In Donohue, 

SJ. (ed.) Reference Soil and Media Diagnostic procedure for the southern region of 

the United States. So. Coop. Series Bulletin 374. Va. Agric. Exp. Station, 

Blacksburg, VA. p. 9-12. 
Sims, J. T. 1989. Comparison of Mehlich 1 and Mehlich 3 extractants for P, K, Ca, Mg, 

Mn, Cu, and Zn in Atlantic Coastal Plain soils. Commun. Soil Sci. Plant Anal. 

20:1707-1726. 
Tran, T. Sen and R.R. Simard. 1993. Mehlich 3 extractable elements, p. 43-49. In M.R. 

Carter (ed.) Soil Sampling and Methods of Analysis. Can. Soc. Soil Sci., Ottawa, 

Ontario. 



18 



Methods for P Analysis, G.M. Pierzynski (ed) 

Tran, T. Sen, M. Giroux, J. Guilbeault, and P. Audesse. 1990. Evaluation of Mehlich 3 

extractant to estimate available P in Quebec soils. Commun. Soil Sci. Plant Anal. 

21:1-28. 
Wolf, A.M. and D.E. Baker. 1985. Comparison of soil test phosphorus by the Olsen, 

Bray PI, Mehlich 1 and Mehlich 3 methods. Commun. Soil Sci. Plant Anal. 16:467- 

484. 



19 



Methods for P Analysis, G.M. Pierzynski (ed) 

Soil Test Phosphorus: Olsen P 
J. Thomas Sims, University of Delaware 



Introduction: 

The "Olsen P" or sodium bicarbonate soil test phosphorus (P) method was developed 
by Sterling R. Olsen and co-workers in 1954 (Olsen et al., 1954) to predict crop response 
to fertilizer P inputs on calcareous soils. It is primarily used in the North Central and 
western United States. The Olsen P method is best suited for calcareous soils, 
particularly those with > 2% calcium carbonate, but has been shown in some research to 
be reasonably effective for acidic soils (Fixen and Grove, 1990). The method is based on 
the use of the HC0 3 ", C0 3 3 and OH in the pH 8.5, 0.5M NaHC0 3 solution to decrease 
the solution concentrations of soluble Ca 2 by precipitation as CaC03 and soluble Al 3+ and 
Fe + by formation of Al and Fe oxyhydroxides, thus increasing P solubility. The increased 
surface negative charges and/or decreased number of sorption sites on Fe and Al oxide 
surfaces at high pH levels also enhance desorption of available P into solution. 

An Olsen P value of 10 mg P/kg is generally considered to be optimum for plant 
growth. This is lower than the critical values used for the Bray and Kurtz P-l, Mehlich 1 
and Mehlich 3 soil tests because the Olsen extractant removes less P from most soils than 
these acidic extractants. Kuo (1996) stated that proper interpretation of Olsen P results for 
soils with diverse properties requires some information on soil P sorption capacity. 
Similarly, Schoenau and Karamanos (1993) cautioned against use of the Olsen test to 
compare P availability in soils with large differences in P chemistry. 

Equipment: 

1. No. 10 (2 mm opening) sieve 

2. Standard 1 g and 2 g stainless steel soil scoops 

3. Automatic extractant dispenser, 25 mL capacity 

4. Extraction vessels, such as 50 mL Erlenmeyer flasks, and filter funnels (9 and 1 1 
cm) and racks 

5. Rotating or reciprocating shaker with a capability of 200 excursions per minute 
(epm) 

6. Whatman No. 42 or No. 2 (or equivalent) filter paper, 9 to 1 1 cm. (Acid resistant 
filter paper may be needed if using an automated method for determining P 
concentration by intensity of color. Bits of filter paper may cause an obstruction in 
the injection valves.) 

Reagents: 

1 . Olsen P Extracting Solution (0.5MNaHCO i , pH 8.5) : Dissolve 420 g commercial- 
grade sodium bicarbonate (NaHC03 ) in distilled water and make to a final volume 
of 10 L. Note that a magnetic stirrer or electric mixer is needed to dissolve the 
NaHC03. Adjust extracting solution pH to 8.5 with 50% sodium hydroxide. 



20 



Methods for P Analysis, G.M. Pierzynski (ed) 

Procedure: 

1. Scoop or weigh 1 g of soil into a 50 mL Erlenmeyer flask, tapping the scoop on 
the funnel or flask to remove all of the soil from the scoop. 

2. Add 20 mL of extracting solution to each flask and shake at 200 or more epm for 
30 minutes at a room temperature at 24 to 27 °C 

3. If it is necessary to obtain a colorless filtrate, add 1 cm (-200 mg) of charcoal 
(DARCO G60, J. T. Baker, Phillipburg, NJ) to each flask. 

4. Filter extracts through Whatman No. 42 filter paper or through a similar grade of 
paper. Refilter if extracts are not clear. 

5. Analyze for P by colorimetry or inductively coupled plasma emission 
spectroscopy using a blank and standards prepared in the Olsen P extracting 
solution. 

Calculations: 

Olsen Extractable P (mg P/kg soil) = 

[Concentration of P in Olsen extract, mg/L ] x [ 0.020 L extract 4- 0.001 kg soil] 

References: 

Fixen, P. E. and J.H. Grove. 1990. Testing soils for phosphorus, p. 141-180. InR.L. 

Westerman (ed.) Soil Testing and Plant Analysis. SSSA, Madison, WI. 
Kuo, S. 1996. Phosphorus, p. 869-919. In D.L. Sparks, (ed.). Methods of Soil Analysis: 

Part 3- Chemical Methods. SSSA, Madison, WI. 
Olsen, S.R., C.V. Cole, F.S. Watanabe, and L.A. Dean. 1954. Estimation of available 

phosphorus in soils by extraction with sodium bicarbonate. USDA Circular 939. 

U.S. Government Printing Office, Washington D.C. 
Schoenau, J.J. and R.E. Karamanos. 1993. Sodium bicarbonate extractable P, K, and N. 

p. 51-58. In M. R. Carter (ed.) Soil Sampling and Methods of Analysis. Can. Soc. 

Soil Sci., Ottawa, Ontario. 



21 



Methods for P Analysis, G.M. Pierzynski (ed) 

A Phosphorus Sorption Index 
J. Thomas Sims, University of Delaware 



Introduction: 

The phosphorus (P) sorption capacity of soils is typically determined by the use of 
batch equilibrium experiments that are used to generate sorption isotherms. These 
isotherms are plots of the amount of P adsorbed from several solutions of known initial 
concentration vs. the P concentration at equilibrium for each solution. For example, Nair 
et al., (1984) proposed, based on an interlaboratory comparison study, a standard 
approach to construct P sorption isotherms, using a soihsolution ratio of 1:25 (w:v), six 
initial P concentrations (as KH2PO4 in a 0.0 1M CaCb matrix), and a 24 h equilibration 
period Results from sorption isotherms can be used to calculate P sorption maxima and P 
bonding energies for soils with different properties and/or as influenced by cultural 
practices, such as crop rotation, tillage, and manuring. 

While useful for agronomic and environmental characterization of the P sorption 
capacity of soils, P sorption isotherms are too time-consuming, complicated, and 
expensive for routine use. To overcome these obstacles Bache and Williams (1971) 
developed a "P Sorption Index" (PSI) that could rapidly determine soil P sorption 
capacity. They evaluated 12 approaches and found that a PSI derived from a single-point 
isotherm (P sorbed from a single solution containing 50 umol P/g soil) was easy to use 
and well correlated with the P sorption capacity of 42 acid and calcareous soils from 
Scotland (r=0.97***)- Other researchers have used the PSI, or modified versions, and 
shown it to be well correlated with soil P sorption capacity determined from complete 
sorption isotherms for soils of widely varying chemical and physical properties 
(Mozaffari and Sims, 1994; Sharpley et al, 1984; Simard et al, 1994). In most cases 
these researchers have maintained the original ratio of added P to soil (1.5 g/kg), but have 
slightly changed the soil: solution ratio, background electrolyte, and/or shaking time. 
Most of these modifications have not affected the correlations between P sorption 
capacity estimated from the PSI and that determined by a full sorption isotherm. The 
procedure described below is based on Bache and Williams (1971). Details on other 
approaches are available in the references cited above. 

Equipment: 

1. Centrifuge and 50 mL polyethylene centrifuge tubes. 

2. Shaker (end-over-end shaker preferred to ensure thorough mixing of soil and 
sorption solution). 

3. Millipore filtration apparatus (0.45-um pore size filters) and vacuum flasks. 

4. 50 mL screw-top test tubes. 

Reagents: 

1. Phosphorus Sorption Solution (75 mg P/L): Dissolve 0.3295 g of monobasic 
potassium phosphate (KH2PO4) in 1 L of deionized H 2 0. Store in refrigerator 
until use. 



22 



Methods for P Analysis, G.M. Pierzynski (ed) 

Procedure: 

1. Weigh 1.00 g of air-dried, sieved (2 mm) soil into a 50 mL centrifuge tube. 

2. Add 20 mL of the 75 mg P/L sorption solution to the centrifuge tube. (Note: This 
provides a ratio of 1.5 g P /kg soil). Add two drops of toluene or chloroform to 
inhibit microbial activity. 

3. Place the tubes in the end-over-end shaker and shake for 18 h at 25+2°C. 

4. Centrifuge the samples at 2000 rpm for 30 minutes. 

5. Using the Millipore filtration apparatus, 0.45-(im filters, and large vacuum flasks, 
filter the centrifugate into 50 mL screw-top test tubes within the flask. 

6. Measure P concentration in the centrifugate colorimetrically or by inductively 
coupled plasma emission spectroscopy (ICP-AES). 

Calculations: 

The PSI has usually been calculated as follows, although some studies have shown 
that expressing PSI directly in mg/kg is acceptable. 

PSI(Lkg l )=-^- 
logC 

where: 

v D uAt o/u n (75 mg P/L -P f )x (0.020 L) 

X = P sorbed (mgP/kg) = —. — — ^ 

(0.00 lkg soil) 

C = P concentration at equilibrium (mg/L), 

and 

Pf = Final P concentration after 18 h equilibration (in mg/L). 

References: 

Bache, B.W., and E.G. Williams. 1971. A phosphate sorption index for soils. J. Soil 

Sci. 22:289-301. 
Mozaffari, P.M., and J.T. Sims. 1994. Phosphorus availability and sorption in an 

Atlantic Coastal Plain watershed dominated by intensive, animal-based agriculture. 

Soil Sci. 157:97-107. 
Nair, P.S., T.J. Logan, A.N. Sharpley, L.E. Sommers, M.A. Tabatabai, and TL. Yuan. 

1984. Interlaboratory comparison of a standardized phosphorus adsorption 

procedure. J. Environ. Qual. 13:591-595. 
Sharpley, A.N., S.J. Smith, B.A. Stewart, and A.C. Mathers. 1984. Forms of phosphorus 

in soils receiving cattle feedlot waste. J. Environ. Qual. 13:211-215. 
Simard, R.R., D. Cluis, G. Gangbazo, and A. Pesant. 1994. Phosphorus sorption and 

desorption indices for soils. Commun. Soil Sci. Plant Anal. 25:1483-1494. 



23 



Methods for P Analysis, G.M. Pierzynski (ed) 

Determination of Water- and/or Dilute Salt-Extractable 
Phosphorus 

M.L. Self-Davis, University of Arkansas 
P.A. Moore, Jr., USDA-ARS, Fayetteville, AR 
B.C. Joern, Purdue University 

Introduction: 

Many methods exist to determine the various forms of soil phosphorus (P). Early 
interests in examining soil P were primarily based on determining the quantity of 
supplemental P needed to adequately meet the needs of crops. The method for using 
distilled water as an extractant to determine P needs of plants was examined in a paper by 
Luscombe et al. (1979). They found a good correlation between the concentration of 
water-extractable P and dry matter yield responses in ryegrass. 

There is now a national focus on examining excessive P buildup in the soil and 
consequent excessive P concentrations in runoff from agricultural land. Many studies 
have examined methods that best correlate soil P levels to concentrations of P in runoff 
(Sharpley, 1995; Pote et al., 1996). The study conducted by Pote et al. (1996) found an 
excellent correlation between water extractable soil test P and dissolved reactive P 
concentrations in runoff. 

One criticism of various other extractants is that they are either more acid or alkaline 
than the soil solution. Therefore, a portion of P extracted is actually of low availability. 
For example, extractants such as Mehlich 3, which contain strong acids, would be 
expected to dissolve calcium phosphates. Also, due to the specific chemical nature of 
many extractants, their use is limited to specific soil types. Using distilled water or 0.01 
MCaCl2 overcomes these criticisms (Pote et al., 1995). 

The following methods are variations of the method described by Olsen and Sommers 
(1982) for determination of water-soluble P in soils. 

Equipment: 

1. Shaker (reciprocating or end-over-end). 

2. Centrifuge. 

3. Centrifuge tubes (40 mL). 

4. Filtration apparatus (0.45 (im pore diameter membrane filter, or Whatman No. 42). 

5. Spectrophotometer with infrared phototube for use at 880 nm. 

6. Acid washed glassware and plastic bottles: graduated cylinders (5 mL to 100 mL), 
volumetric flasks (100 mL, 500 mL, and 1000 mL), storage bottles, pipets, dropper 
bottles, and test tubes or flasks for reading sample absorbance. 

Reagents: 

1. Concentrated hydrochloric acid (HC1). 

2. Reagents used for ascorbic acid technique for P determination, Murphy and Riley 
(1962). 

3. M calcium chloride (CaCh ). 

4. Chloroform. 



24 



Methods for P Analysis, G.M. Pierzynski (ed) 

Extraction Procedure - Deionized Water: 

Weigh out 2 g of soil (dried in a forced-draft oven at 60°C for 48 hours, sieved 
through a 2-mm mesh sieve) into a 40 mL centrifuge tube. Add 20 mL of distilled water 
and shake for one hour. Centrifuge at 6,000 rpm for 10 minutes. Filter the solution 
through a 0.45 (im membrane filter. Acidify to pH 2.0 with HC1 to prevent precipitation 
of phosphate compounds (approximately 2 days of concentrated HC1). Freeze the 
sample if it is not going to be analyzed that day. Previous articles have noted that 
hydrolysis of condensed phosphates can occur when the solution is acidified (Lee et al., 
1965). Also, at this pH level, there is the possibility of flocculation of organics. 
However, it is vital to ensure that the P remains in solution, therefore, we consider the 
negative effects of acidification minimal. 

Extraction Procedure - 0.01M CaCl 2 : 

Weigh out 1 g of dry soil into a 40 mL centrifuge tube. Add 25 mL of 0.01 M CaCl 2 
(you can add 2 drops of chloroform to inhibit microbial growth if desired) and shake for 
one hour on a reciprocating shaker. Centrifuge at 4000 rpm for 10 minutes. Filter 
solution through Whatman No. 42 filter paper. 

Analysis: 

For determining water or dilute salt extractable P in soil, any spectrophotometer with 
an infrared phototube for use at 660 or 882 nm can be used. Also, samples can be 
analyzed by inductively coupled plasma-atomic emission spectrometry (ICP-AES), 
which will measure total dissolved P. 

Calculations: 

Water- or Dilute salt-extractable P (mg P/kg soil) = 

[Concentration of P in extract, mg/L] x [Volume of extractant, L 4- mass of soil, kg] 

Comments: 

It should be mentioned that some studies have shown that concentrations of P in CaCh. 
extracts can be one-third to one-half that of water extracts (Olsen and Watanabe, 1970; 
Soltanpour et al., 1974). Concentrations of Ca were less in the water extracts, as 
compared with the CaCh extracts, which resulted in higher P concentrations in the water 
extracts. Higher concentrations of Ca in the extracting solution may precipitate calcium 
phosphate, lowering the P levels in solution. 

References: 

Lee, G.R., N.L. Clesceri, and G.P. Fitzgerald. 1965. Studies on the analysis of phosphates 

in algal cultures. Internat. J. Air Water Poll. 9:715-722. 
Luscombe, P.C., J.K. Syers, and P.E.H. Gregg. 1979. Water extraction as a soil testing 

procedure for phosphate. Commun. Soil Sci. Plant Anal. 10:1361-1369. 
Murphy, J., and J. P. Riley. 1962. A modified single solution method for the 

determination of phosphate in natural waters. Anal. Chem. Acta 27:31-36. 
Olsen, S.R., and L.E. Sommers. 1982. Phosphorus. P. 403-430 In A.L. Page et al. (ed.) 

Methods of soil analysis. Part 2. 2nd ed. Agronomy Monogr. 9. ASA and SSSA, 

Madison, WI. 



25 



Methods for P Analysis, G.M. Pierzynski (ed) 

Olsen, S.R., and F.S. Watanabe. 1970. Diffusive supply of phosphorus in relation to soil 

textural variations. Soil Sci. 110:318-327. 
Pote, D.H., T.C. Daniel, P.A. Moore, Jr., A.N. Sharpley, D.R. Edwards, and D.J. Nichols. 

1995. Phosphorus: relating soil tests to runoff concentrations across five soil series. 
Agronomy Abstracts, p. 294, Am. Soc. Agron., Madison, WI. 

Pote, D.H., T.C. Daniel, A.N. Sharpley, P.A. Moore, Jr., D.R. Edwards, and D.J. Nichols. 

1996. Relating extractable soil phosphorus to phosphorus losses in runoff. Soil Sci. 
Soc. Am J. 60:855-59. 

Sharpley, A.N. 1995. Dependence of runoff phosphorus on extractable soil phosphorus. 

J. Environ. Qual. 24:920-926. 
Soltanpour, P.N., F. Adams, and A.C. Bennett. 1974. Soil phosphorus availability as 

measured by displaced soil solutions, calcium chloride extracts, dilute-acid extracts, 

and labile phosphorus. Soil Sci. Soc. Am. Proc. 38:225-228. 



26 



Methods for P Analysis, G.M. Pierzynski (ed) 

Phosphorus Extraction with Iron Oxide-Impregnated Filter 
Paper (Pi test) 

W.J. Chardon, DLO Research Institute for Agrobiology and Soil Fertility, 
The Netherlands 

Introduction: 

The availability of phosphorus (P) in soil or surface water for biota (e.g. plants or 
algae) has been studied extensively, and numerous tests for available P have been 
developed and used. These tests can roughly be divided into four categories: (1) shaking 
with acid solutions which dissolve P compounds or with (buffered) alkaline solutions 
which displace P from the soil; (2) measuring exchangeable P, using P; (3) shaking with 
dilute salt solutions or water, which simulate the soil solution, and (4) as (3), with a sink 
added, acting more or less analogous to the withdrawing behavior of a plant root. 

The use of resin beads as a sink for P was introduced by Amer et al. (1955). Stronger 
sinks for P were developed by Hsu and Rich (1960) and by Robarge and Corey (1979) 
who affixed hydroxy- Al to a cation exchange resin. Since the use of these resins is 
laborious, it has not developed into a practical method (T.C. Daniel, pers. 
communication). Iron (hydr)oxide impregnated filter paper (FeO paper, also known as Pi 
paper or HFO paper) was initially developed for soil chemical studies in the late '70s. 
Later, it was introduced for plant availability studies as a simpler alternative for resin 
beads. A water extraction procedure is used for fertilizer recommendations in the 
Netherlands. In tropical soils this method often results in very low amounts of extracted 
P, causing analytical problems. Therefore, FeO paper was added as a sink during the 
extraction. However, since the use of water as an extractant allowed soil dispersion with 
resulting contamination of the FeO paper with soil particles, 0.01 M CaCb. was chosen as 
an alternative for water. Although the description of the preparation of the FeO paper and 
its application was only published in an internal report (Sissingh, 1983), its use became 
widespread. The application for plant availability studies was reviewed by Menon et al. 
(1990, 1997), and the use for water-quality studies was described by Sharpley et al. 
(1995). For long-term desorption studies, an alternative method was developed using a 
FeO- suspension in a dialysis bag (Lookman et al., 1995). The present paper is mainly 
based on Chardon et al. (1996), in which studies on the various aspects of both 
preparation and use of the FeO paper are reviewed in a historical perspective. 

Principle of the method: 

Filter paper is covered with a precipitate of amorphous iron(hydr)oxides (FeO). When 
a soil is shaken in CaCb to which a strip of this FeO paper is added, P will first desorb 
from the soil, then adsorb onto the FeO- strip and new P will desorb from the soil. During 
shaking, the desorbable fraction of soil P will thus be (partly) depleted. During shaking 
the strip is protected against erosion by soil particles via a polyethylene screen. After 
shaking the strip is taken out and adhering soil particles are removed by rinsing with 
distilled water using an air-brush. The FeO on the paper with the P adsorbed onto it is 
dissolved in H 2 SO4 and P is determined in the acidic solution. 



27 



Methods for P Analysis, G.M. Pierzynski (ed) 

Equipment: 

1. 15-cm discs of ash-free, hard filter paper (e.g. Schleicher & Schuell 589 red ribon 
or Whatman No. 50) 

2. Tweezers 

3. Immersing baths 

4. Polyethylene shaking bottles (100 mL) 

5. Polyethylene screen (925 urn openings) 

6. Shaking apparatus, end-over-end 

7. Air brush 

Reagents: 

1. Acidified FeC13 solution: completely dissolve 100 g FeC13 in 1 10 mL 
concentrated HC1 and dilute with distilled water to 1 L. 

2. 5 % NH40H: dilute 200 mL NH40H (25%) to 1 L with distilled water. 

3. 0.01 M CaC12: stock solution 0.1 M: dissolve 14.7 g CaC12.2 H20 in distilled 
water and dilute to 1. L; reagent 0.01 M: dilute the stock solution tenfold with 
distilled water. 

4. 0. 1 M H2S04: stock solution 2.5 M: add 140 mL of concentrated H2S04 to 750 
mL distilled water, cool and dilute with distilled water to 1 L; reagent 0.1 M, dilute 
40 mL of 2.5 M H2S04 to 1 L with distilled water. 

5. Distilled/deionized water 

Procedures: 

Preparation ofFeO paper 

1. Immerse the filter paper in acidified FeC13, using tweezers, for at least 5 minutes. 

2. Let the paper drip dry at room temperature for 1 h. 

3. Pull the paper rapidly and uninterrupted through a bath containing 2.7 M NH40H 
to neutralize the FeC13 and produce amorphous iron (hydr)oxide (ferrihydrite, 
denoted as FeO). 

4. Rinse the paper with distilled water to remove adhering particles of FeO. 

5. After air drying, cut the paper into strips with a (reactive) surface of40 cm2 
(generally 2 by 10 cm). 

Shaking soil suspension with FeO strip added 

1. Add 40 mL 0.01 M CaC12 to 1 g of soil in a 100 mL bottle; add one strip protected 
by polyethylene screen, in a fixed position, at room temperature. 

2. Shake on a reciprocating shaker at a speed of 130 excursions/min, or at 4 rpm end- 
over-end, for 16 h. 

3. Take out the strip, thoroughly rinse with distilled water to remove adhering soil 
particles using an air brush, and remove adhering water. 

Determination ofP extracted by FeO paper 

Dissolve the FeO with adsorbed P by shaking 1 h in 40 mL 0.1 MH2SO4 and 
determine P in the acidic extract with colorimetry or by inductively coupled plasma 
spectrophotometry. 



28 



Methods for P Analysis, G.M. Pierzynski (ed) 

Calculations: 

The FeO-extractable P content of a soil, also called Pi- value, is expressed as mg 
P/kg soil, and can be calculated as: 

C.V 



P value 



W 



where: 



C p = P concentration in H2SO4, mg/L, 
V = volume of H 2 SO4 , L, 
W = mass of soil used, kg. 

Comments: 

The method described above can be used as a standard method to estimate soil plant- 
available P content. In case total desorbable P is studied one can use more FeO-strips 
during shaking, increase the shaking time, or the amount of FeO on a strip by using a 
higher concentration of FeCL (Chardon et al., 1996). When long-term desorption kinetics 
is studied the shaking time can be increased, the paper can be refreshed e.g. daily 
(Sharpley, 1996), or the technique with an FeO-filled dialysis membrane can be used 
(Freese et al., 1995, Lookman et al., 1995). Myers et al. (1997) described the use of 5.5 
cm diameter filter paper circles, which eliminates the need for cutting strips. 

As discussed in detail in Chardon et al. (1996) soil particles adhering to the strip 
when the FeO on the strip is dissolved in H2SO4 may give erroneous results, since P from 
the soil particles can also dissolve in the acid as if it was desorbed. The use of a nylon 
screen around the strip during shaking and an air-brush after shaking to clean the strip 
(Whelan et al., 1994) will strongly reduce this risk. Since temperature influences P 
desorption it is recommended to perform the procedure at a constant temperature in order 
to get reproducible results. 

References: 

Amer, F., D.R. Bouldin, C.A. Black, and F.R. Duke. 1955. Characterization of soil 

phosphorus by anion exchange resin adsorption and P 32 -equilibration. Plant Soil 6: 

391-408. 
Chardon, W.J., R.G. Menon, and S.H. Chien. 1996. Iron oxide-impregnated filter paper 

(Pi test): a review of its development and methodological research. Nutr. Cycl. in 

Agroecosyst. 46:41-51. 
Freese, D., R. Lookman, R. Merckx, and W.H. van Riemsdijk. 1995. New method for 

assessment of long-term phosphate desorption from soils. Soil Sci. Soc. Am. J. 

59:1295-300. 
Hsu, P.S. and C.I. Rich. 1960. Aluminum fixation in a synthetic cation exchanger. Soil 

Sci. Soc. Am. Proc. 24: 21-25. 
Lookman, R., D. Freese, R. Merckx, K. Vlassak, and W.H. van Riemsdijk. 1995. Long- 
term kinetics of phosphate release from soil. Environ. Sci. Technol. 29: 1569-1575. 
Menon, R.G., S.H. Chien, and L.L. Hammond. 1990. Development and evaluation of 

the Pi soil test for plant-available phosphorus. Commun. Soil Sci. PL Anal. 21: 1131- 

1150. 



29 



Methods for P Analysis, G.M. Pierzynski (ed) 

Menon, R.G., S.H. Chien, and W.J. Chardon. 1997. Iron hydroxide-impregnated filter 

paper (Pi test): II. A review of its application. Nutr. Cycl. in Agroecosyst. 47:7-18. 
Myers, R.G., G.M. Pierzynski, and S.J. Thien. 1997. Iron oxide sink method for 

extracting soil phosphorus paper preparation and use. Soil Sci. Soc. Am. J. 61:1400- 

1407. 
Robarge,W.P. and R.B. Corey. 1979. Adsorption of phosphate by hydroxy- aluminum 

species on a cation exchange resin. Soil Sci. Soc. Am. J. 43: 481-487. 
Sharpley, A.N., J.S. Robinson, and S.J. Smith. 1995. Bioavailable phosphorus dynamics 

in agricultural soils and effects on water quality. Geoderma 67: 1-15. 
Sharpley, A.N. 1996. Availability of residual phosphorus in manured soils. Soil Sci. 

Soc. Am. J. 60:1459-1466. 
Sissingh, HA. 1983. Estimation of plant-available phosphates in tropical soils. A new 

analytical technique. Haren, Netherlands, Inst Soil Fertility Research. 
Whelan, B.R., K. Wittwer, S.P. Roe, N.J. Barrow, and R.G.V. Bramley. 1994. An air 

brush and water to clean the iron oxide strips in the Pi soil test for phosphorus. Trans. 

15th Int. Congress Soil Sci., Acapulco, Mexico, 1994, vol 5b pp 166-167. 



30 



Methods for P Analysis, G.M. Pierzynski (ed) 

Determination of the Degree of Phosphate Saturation In Non- 
Calcareous Soils 

O.F. Schoumans, Winand Staring Centre for Integrated Land, Soil and 
Water Research, The Netherlands 

Introduction: 

The transport of phosphorus (P) by leaching, erosion and surface runoff from 
agricultural soils can contribute to the eutrophication of surface waters. In flat areas with 
shallow groundwater tables, like many areas in the Netherlands, leaching can be an 
important transport pathway. In order to quantify the eutrophication risk of agricultural 
land in areas with intensive livestock production in the Netherlands (non-calcareous 
sandy soils), the degree of P saturation of soils has been introduced as a simple index 
(Breeuwsma and Schoumans, 1987; Breeuwsma et al., 1995). The degree of P saturation 
(DPS) is defined as the ratio between the amount of phosphate accumulated in soils to a 
critical depth (P ac t) and the maximum phosphate sorption capacity (PSC) of the soil to 
that depth. The relationship is described by: 

DPS =^^* 100 
PSC 

Eq.(l) 
where 

DPS = degree of phosphate saturation (%), 

Pact = actual amount of sorbed phosphate to the critical depth (mmol/kg), and 

PSC = maximum phosphate sorption capacity to critical depth (mmol/kg) 

In the Netherlands the mean highest groundwater level (MHW) is used as a critical 
depth. The phosphate sorption capacity of soils depends on soil characteristics (e.g. 
aluminium, iron, clay, lime and organic matter). In acid to neutral soils fixation of P 
mainly takes place with reactive forms of Fe and Al (as hydroxides and Al and Fe bound 
to the organic matter). These reactive forms of Fe and Al can be extracted from soil 
samples (Beek, 1978; Schwertmann, 1964) by shaking at a 1:20 weight to volume ratio 
with a solution of oxalic acid and ammonium oxalate having a nearly constant pH of 3. 
The phosphate sorption capacity of non-calcareous sandy soils can be assessed by 
(Schoumans et al., 1986; Van der Zee, 1988): 



P5C = £0.5(A/„+FO,*P^-*A 

Eq. (2) 



i=i 



where 



Al ox = oxalate extractable aluminium of soil layer i (mmol/kg), 
Fe ox = oxalate extractable iron of soil layer i (mmol/kg), 
pd,i = dry bulk density of soil layer i (kg/m), 
Li = thickness of soil layer i (m), and 
n = amount of observed layers. 



31 



Methods for P Analysis, G.M. Pierzynski (ed) 

The amount of P which is bound to the reactive amount of Al and Fe comes into 
solution with the oxalate extraction. Therefore, the actual amount of sorbed P can be 
calculated by means of: 

n 

P = V P * n * 1 

1 act L^ 1 oxj Hd,f L -'i 

Eq. (3) 
where 

Pox = oxalate extractable P of soil layer i (mmol/kg). 

If the dry bulk densities of the observed layers (from the soil surface to the reference 
depth) are identical, or a soil sample has been taken over the complete depth (on volume 
basis), the degree of P saturation can be calculated by the mean contents of P ox , Al ox and 
Fe ox (in mmol/kg) over the observed depth: 

P 

DPS = ^ * 100 

0.5(AZ„+FO 

Eq. (4) 

Based on desorption characteristics of non-calcareous sandy soils, Van der Zee et al. 
(1990) have show that at a degree of P saturation of 25% the P concentration in pore 
water will become higher than 0.1 mg/L ortho-P at the long term (after redistribution of 
the P front in the soil). In the Netherlands this concentration is used as a target level at the 
mean highest water table. 

A disadvantage of the definition of the phosphate saturation degree is that this 
parameter depends on the phosphate sorption capacity of the soil (Equation 1), which 
varies from layer to layer and which is in most situations assessed (e.g., for non- 
calcareous sandy soils by means of 0.5 (Al ox + Fe ox ) ). In order to omit this assessment of 
the phosphate sorption capacity also an independent P saturation index (PSI) can be used: 

P 

PSI = 



AL+Fe ox 

Eq. (5) 

Reagents: 

1. Extraction solution (pH = 3) . Dissolve 16.2 g of ammonium oxalate monohydrate. 
(COONH 4 ) 2 .H 2 and 10.8 g of oxalic acid dihydrate, (COOH 2 ).2H 2 in water in a 
1000 mL volumetric flask. The pH of this solution must be 3.0 + 0.1. 

2. Hydrochloric acid. 1 M. Dilute 83 mL of concentrated hydrochloric acid, HC1 (p= 
1.19 glcm ), with water to volume of 1000 mL. 

3. Hydrochloric acid. 0.01 M. Dilute 10 mL of \M hydrochloric acid with water to 
volume of 1000 mL. 

4. Standard Fe solution . 1000 mg/L 



32 



Methods for P Analysis, G.M. Pierzynski (ed) 

5. Standard Al solution . 1000 mg/L 

6. Standard P solution. 500 mg/L. Dissolve 2. 1950 g of potassium dihydrogen 
phosphate (KH2PO4) in water in a volumetric flask of 1000 mL and dilute to 1000 
mL with water. 

Procedure: 

The method, which is described below, is a summary of the Dutch norm (NEN 5776). 
Weigh 2.5 (± 0.01) g of air-dry soil (< 2 mm) in a dry, 100 mL polyethene bottle. Add 
with a dispenser 50 mL of the oxalate extraction solution (1) and close the bottle. Prepare 
two blanks and take three reference samples. Shake at 180 excursions/min on a 
reciprocating shaker for 2 hours in a darkened conditioned room at constant temperature 
(20 °C). Filter the extracts through a fine filter paper (high quality). Discard the first three 
mL of the filtrate and collect the remainder in a 100 mL polyethene bottle. Pipet 10 mL 
of the soil extracts in flasks. Add 40 mL of 0.01 M HCl-solution (3) and mix. Measure 
the concentration of P, Al and Fe within one week with the ICP-AES. 

Pipet 0, 2.5, 10.0, 25.0 and 50.0 mL of each standard element solution ((4), (5) and 
(6)) in a volumetric flask of 1000 mL Add 10 mL of (1M HC1) and 200 mL of 
(extraction solution) and mix. Dilute to 1000 mL with water. This standard series 
contains 0, 1.25, 5.0, 12.5 and 25.0 mg/L P and 0, 2.5, 10.0, 25.0 and 50.0 mg/L Al and 
Fe. 

Comments: 

The extraction should be performed in dark because the extraction solution (1) 
partially reduces the poorly soluble iron(lll) ions to the much more soluble iron(ll) ions 
and light influences the reducing action of oxalic acid. 

The soil filtrates should be stored in a refrigerator if they are not used directly for 
analysis. 

Calculation: 

(a-b)* 0.05 



P.. = 



m* 30.97 



(a-b)*0.05 

Fe os = , and 

m* 55.85 



Al 



(a-b)* 0.05 
m* 26.98 



where: 



P ox , Fe os , Al ox = content of P, Fe and Al of the air-dry soil sample in mmol/kg 
a = concentration of P, Fe, Al in the soil extraction solution in 

mg/L 



33 



Methods for P Analysis, G.M. Pierzynski (ed) 

b = concentration of P, Fe, Al in the blank extraction solution in 

mg/L 
m = air-dry soil sample weight in grams. 

P 

PSI = 



AL + Fe ox 



DPS= 200 PSI 



Comments: 



Since the calculation of the results of soil analysis are generally expressed on an 
"oven-dry" basis, the moisture content of "air-dry" soil should be determined shortly 
before soil analysis and the appropriate correction made. 



References: 

Breeuwsma, A. and O.F. Schoumans, 1987. Forecasting phosphate leaching on a 
regional scale. In: Proceedings of the International Conference on Vulnerability of 
soil and groundwater to pollutants on March 30 to April 3, 1987 at Noordwijk aan 
Zee. (Eds. W. van Duijvenboode and H.G. van Waegeningh). The Hague: TNO 
Committee on Hydrological Research, The Hague, Proceedings and Information No. 
38. 

Breeuwsma, A., J.G.A. Reijerink, O.F. Schoumans, 1995. Impact of manure on 

accumulation and leaching of phosphate in areas of intensive livestock farming: IN: 
Animal Waste and the Land- Water Interface (Ed. K. Steele). Lewis publishers, New 
York, pp. 239-249. 

Beek, J., 1978. Phosphate retention by soil in relation to waste disposal. Ph. D. Thesis, 
Wageningen, The Netherlands. 

NEN 5776, Soil. Determination of iron, aluminum and phosphorus in an ammonium 
oxalate-oxalic acid extract for estimation of the saturation with phosphate. 
Netherlands Normalization Institute (NNI), Delft, The Netherlands. 

Schwertmann, D. 1964, Differentierung der Eisenoxide des Badens durch 

photochemische Extraction mit saurer Ammoniumoxalaat-Losung. Z.Pfl. Ern. 
Duengung 105. 194-202 

Schoumans, O.F., W. de Vries and A. Breeuwsma, 1986. A phosphate transport model 
for application on regional scale. Winand Staring Centre for Integrated Land, Soil and 
Water Research, Wageningen, The Netherlands, rapport 1951 (in Soil Survey series). 

van der Zee, S.E.A.T.M., 1988. Transport of reactive contaminants in hetereogeneous 
soil systems. Ph. D. Thesis, Agricultural University, Wageningen, The Netherlands. 

van der Zee, S.E.A.T.M., W.H. van Riemsdijk, and F.A.M. de Haan, 1990. Protocol 
phosphate saturated soils. Department of soil science and plant nutrition, Agricultural 
University, Wageningen, The Netherlands. 



34 



Methods for P Analysis, G.M. Pierzynski (ed) 

Phosphorus Sorption Isotherm Determination 

D.A. Graetz, University of Florida 
V.D. Nair, University of Florida 

Introduction: 

Phosphorus (P) retention by soils is an important parameter for understanding soil 
fertility problems, as well as for determining the environmental fate of P. The P 
adsorption capacity of a soil or sediment is generally determined by batch-type 
experiments in which soils or sediments are equilibrated with solutions varying in initial 
concentrations of P. Equations such as the Langmuir, Freundlich and Tempkin models 
have been used to describe the relationship between the amount of P adsorbed to the P in 
solution at equilibrium (Berkheiser et al., 1980; Nair et al., 1984). 

Advantages of the batch technique include: the soil and solution are easily separated, a 
large volume of solution is available for analysis, and the methodology can be easily 
adapted as a routine laboratory procedure. Disadvantages include difficulties in 
measuring the kinetics of the sorption reaction and optimizing the mixing of solution and 
soil without particle breakdown (Burgoa et al. 1990). Despite the disadvantages, the 
batch technique has been, and still is, widely used to describe P sorption in soils and 
sediments. 

Nair et al. (1984) noted that P sorption varies with soil/solution ratio, ionic strength 
and cation species of the supporting electrolyte, time of equilibration, range of initial P 
concentrations, volume of soil suspension to head space volume in the equilibration tube, 
rate and type of shaking, and type and extent of solid/solution separation after 
equilibration. Although most researchers use a similar basic procedure for measuring P 
adsorption, there is considerable variation observed among studies with regard to the 
above parameters. This variation often makes comparisons of results among studies 
difficult. Thus, Nair et al. (1984) proposed a standard P adsorption procedure that would 
produce consistent results over a wide range of soils. This procedure was evaluated, 
revised, tested among laboratories and was eventually proposed as a standardized P 
adsorption procedure. This procedure as described below is proposed as the standard 
procedure recommended by the SERA-IEG 17 group. 

Equipment: 

1. Shaker: End-over-end type 

2. Filter Apparatus: Vacuum filter system using 0.45 or 0.2 (im filters 

3. Equilibration tubes: 50 mL or other size to provide at least 50% head space 

4. Spectrophotometer: Manual or automated system capable of measuring at 880 nm 

Reagents: 

1 . Electrolyte: 0.0 1 M CaCl 2 , unbuffered 

2. Microbial inhibitor: Chloroform 

3. Inorganic P solutions: Selected concentrations as KH2PO4 or NaH 2 P04 (in 0.01 M 
CaCb containing: 20 g/L chloroform) 



35 



Methods for P Analysis, G.M. Pierzynski (ed) 

Procedure: 

1. Air-dry soil samples and screen through a 2 mm sieve to remove roots and other 
debris. 

2. Add 0.5 to 1.0 g air-dried soil to a 50 mL equilibration tube. 

3. Add sufficient 0.01 M CaCl 2 solution containing 0, 0.2, 0.5, 1, 5, and 10 mg P/L as 
KH2PO4 or NaH 2 P04, to produce a soihsolution ratio of 1:25. The range of P 
values could vary from to 100 mg P/L (0, 0.01, 0.1, 5, 10, 25, 50 and 100 mg 
P/L) and the soil/solution ratio could be as low as 1:10 depending on the sorbing 
capacity and the P concentrations of the soils in the study. 

4. Place equilibration tubes on a mechanical shaker for 24 h at 25 + 1 C. 

5. Allow the soil suspension to settle for an hour and filter the supernatant through a 
0.45 (im membrane filter. 

6. Analyze the filtrate for soluble reactive P (SRP) on a spectrophotometer at a 
wavelength of 880 nm. 

Calculations and Recommended Presentation of Results: 

Two of the often used isotherms are the Langmuir and the Freundlich isotherms; the 
Langmuir having an advantage over the Freundlich in that it provides valuable 
information on the P sorption maximum, S m ax and a constant k, related to the P bonding 
energy. 

The Langmuir equation 

The linearized Langmuir adsorption equation is: 

C 1 C 

- + - 



S kS„„ s 



max 



where: 



S = S' + S„, the total amount of P retained, mg/kg 

S' = P retained by the solid phase, mg/kg 

S = P originally sorbed on the solid phase (previously adsorbed P), mg/kg 
C = concentration of P after 24 h equilibration, mg/L 
Smax = P sorption maximum, mg/kg, and 
k = a constant related to the bonding energy, L/mg P. 

The Freundlich equation 

The linear form is: log S = log K + n log C 
where: 

K is the adsorption constant, expressed as mg P/kg, 
n is a constant expressed as L/kg, and 
C and S are as defined previously. 

A plot of log S against log C will give a straight line with log K as the intercept, and n 
as the slope. 



36 



Methods for P Analysis, G.M. Pierzynski (ed) 

Previously adsorbed P (also referred to as native sorbed P) 

Adsorption data should be corrected for previously adsorbed P (S ). For the 
calculation of previously sorbed P, Nair et al. (1984) used isotopically exchangeable P 
(Holford et al., 1974) prior to calculations by the Langmuir, Freundlich and Tempkin 
procedures. Other procedures used to calculate the previously adsorbed P include 
oxalate-extractable P (Freese et al., 1992; Yuan and Lavkulich, 1994), anion-impregnated 
membrane (AEM) technology (Cooperband and Logan, 1994) and using the least squares 
fit method (Graetz and Nair, 1995; Nair et al, 1998; Reddy et al, 1998). Sallade and 
Sims (1997) used Mehlich 1 extractable P as a measure of previously sorbed P. 

Investigations by Villapando (1997) have indicated a good agreement among native 
sorbed P values estimated by the least squares fit method, oxalate extractions, and the 
AEM technology. At this point, it appears that selection of the method for determination 
of native sorbed P would depend on the nature of the soils in the study and 
reproducibility of the results. 

The procedure for calculation of S using the least square fit method is based on the 
linear relationship between S and C at low equilibrium P concentrations. The 
relationship can be described by 

S = KC - S„ 
where 

K = the linear adsorption coefficient, and 
all other parameters are as defined earlier 
(Note: It is recommended that the linear portion of the isotherm has an r value 0.95 or 
better). 

Equilibrium P Concentration 

The "equilibrium P concentration at zero sorption" (EPCo) represents the P 
concentration maintained in a solution by a solid phase (soil or sediment) when the rates 
of P adsorption and desorption are the same (Pierzynski et al., 1994). Values for EPCo 
can be determined graphically from isotherm plots of P sorbed vs. P in solution at 
equilibrium. From the calculations given above, EPCo is the value of C when S = 0. 

Comments: 

The above procedure was developed to provide a standardized procedure with a fixed 
set of conditions that could be followed rigorously by any laboratory. The procedure 
uses a low and narrow range of dissolved inorganic P concentrations because these are 
the concentrations likely to be encountered in natural systems and because higher 
concentrations may result in precipitation of P solid phases. However, higher 
concentrations of P (up to 100 mg/L) and/or lower soihsolution ratios (1:10) have been 
used for isotherm determinations on soils and sediments (Mozaffari and Sims, 1994; 
Sallade and Sims, 1997; Nair et al, 1998; Reddy et al, 1998). A 0.01 MKC1 solution 
may be used as the background electrolyte to avoid precipitation of Ca in neutral and 
alkaline soils. 

Toluene and chloroform have been shown to increase the dissolved P concentration in 
the supernatant, apparently due to lysis of microbial cells, and thus, some researchers do 
not try to inhibit microbial growth (Reddy et al., 1998). 



37 



Methods for P Analysis, G.M. Pierzynski (ed) 

Most adsorption studies are conducted under aerobic conditions, however, with certain 
studies it is more appropriate to use anaerobic conditions, as they more closely represent 
the natural environments of the soils or sediments. Reddy et al. (1998) preincubated 
sediment/soil samples in the dark at 25°C under a N2 atmosphere, to create anaerobic 
conditions. Adsorption experiments were then conducted, performing all equilibrations 
and extractions in an Ch-free atmosphere. 

References: 

Berkheiser, V.E., J.J. Street, P.S.C. Rao, and T.L. Yuan. 1980. Partitioning of inorganic 

orthophosphate in soil-water systems. CRC Critical Reviews in Environmental 

Control. 179-224. 
Burgoa, B., R.S. Mansell, and D. Rhue. 1990. Effects of solids concentrations upon non- 
equilibrium phosphorus sorption in aqueous soil suspensions. Soil and Crop Sci. Soc. 

Fla. Proc. 49:60-66. 
Cooperband, L.R., and Logan. 1994. Measuring in situ changes in labile soil phosphorus 

with anion-exchange membranes. Soil Sci. Soc. Am. J. 58:105-114. 
Freese, D., S.EA.T.M. van der Zee, and W.H. van Riemsdijk. 1992. Comparison of 

different models for phosphate sorption as a function of the iron and aluminum oxides 

of soils. J. Soil Sci. 43:729-738. 
Graetz, DA., andV.D. Nair. 1995. Fate of phosphorus in Florida Spodosols 

contaminated with cattle manure. Ecol. Eng. 5:163-181. 
Holford, I.C.R, R.W.M. Wedderburn, and G.E.G. Mattingly. 1974. A Langmuir two 

surface equation as a model for phosphate adsorption by soils. J. Soil Sci. 25:242- 

255. 
Mozaffari, M. and J.T. Sims. 1994. Phosphorus availability and sorption in Atlantic 

Coastal Plain watershed dominated by animal-based agriculture. Soil Sci. 157:97-107. 
Nair. P.S., T.J. Logan, A.N. Sharpley, L.E. Sommers, MA. Tabatabai, and T.L. Yuan. 

1984. Interlaboratory comparison of a standardized phosphorus adsorption procedure. 

J. Environ. Qual. 13:591-595. 
Nair, V.D., D.A. Graetz, and K.R. Reddy. 1998. Dairy manure influences on phosphorus 

retention capacity of Spodosols. J. Environ. Qual. 27:522-527. 
Pierzynski, G.M., J.T. Sims and G.F.Vance. 1994. Soils and Environmental Quality. 

Lewis Publishers, Boca Raton, FL 
Sallade, Y.E., and J.T. Sims. 1997. Phosphorus transformations in the sediments of 

Delaware's agricultural drainageways: I. Phosphorus forms and sorption. J. Environ. 

Qual. 26:1571-1579. 
Reddy, K.R., G.A. O'Connor, and P.M. Gale. 1998. Phosphorus sorption capacities of 

wetland soils and stream sediments impacted by dairy effluent. J. Environ. Qual. 

27:438-447. 
Villapando, R.R. 1997. Reactivity of inorganic phosphorus in the spodic horizon. Ph.D. 

Dissertation, University of Florida, Gainesville, Florida. (Diss. Abstr. Vol. 58. No. 

7B). 258pp. 
Yuan, G., and L.M. Lavkulich. 1994. Phosphate sorption in relation to extractable iron 

and aluminum in Spodosols. Soil Sci. Soc. Am. J. 58:343-346. 



38 



Methods for P Analysis, G.M. Pierzynski (ed) 

Bioavailable Phosphorus in Soil 

Andrew Sharpley, USDA-ARS, University Park, PA 

Introduction: 

Biologically available P (BAP) has been operationally defined as "..the amount of 
inorganic P, a P-deficient algal population can utilize over a period of 24 h or longer" 
(Sonzogni et al., 1982). The amount of P in soil, sediment, and water that is potentially 
available for algal uptake (bioavailable P) can be quantified by algal assays, which 
require up to 100-d incubations (Miller et al., 1978). Thus, more rapid chemical 
extractions, such as those using NaOH (Butkus, et al., 1988; Dorich et al., 1980), NH 4 F 
(Porcella et al., 1970), ion exchange resin (Huettl et al., 1979) and citrate-dithionite- 
bicarbonate (Logan et al., 1979), have been used routinely to estimate bioavailable P. 
The weaker extractants (NH4F and NaOH) and short-term resin extractions may represent 
P that could be utilized by algae in the photic zone of lakes under aerobic conditions. In 
contrast, the more severe extractants (citrate-dithionite-bicarbonate) represent P that may 
become bioavailable under the reducing conditions found in the anoxic hypolimnion of 
stratified lakes. 

Sharpley et al. (1991) showed that when using a wide solution:soil ratio (500:1), 0.1 M 
NaOH extractable P (NaOH-P) was closely related to the growth of several algal species. 
However, the complexity of algal assay and chemical extraction methods often limits 
their use by soil testing laboratories. For example, long assay incubation (7 to 100 d) and 
chemical extraction times (> 16 hr), as well as large solution volumes (> 500 mL) are 
particularly inconvenient. As the amount of P extracted depends on ionic strength, 
cationic species, pH, and volume of the extractant used (Hope and Syers, 1976; Sharpley 
et al., 1981), these limitations will be difficult to overcome. Questions also have been 
raised as to the validity of relating the form or availability of P extracted by chemical 
solutions to P bioavailability in the aquatic environment. As a result, P sink approaches 
have been developed to estimate BAP in soil, sediment, and water. 

P-Sink Approaches: 

The concept of exposing the soil to a P-sink has merit toward the goal of assessing 
soil, sediment, and water BAP (i.e., available to plants and algae) for both agronomic and 
environmental goals. Presumably, this would allow only P that was able to respond to 
such a sink to be measured, which is analogous to a root acting as a sink in the soil or to 
the concentration gradient that exists when a small quantity of sediment is placed in a 
large volume of water. The analogy of a root is not entirely accurate because root 
exudates and mycorrhizae fungi can alter P availability in the rhizosphere such that the 
root does not behave as a pure sink. Still, P-sinks are likely the closest manifestation of 
the root environment that are available. Some authors assume that the sink maintains 
extremely low P concentrations in the aqueous media employed and can be considered an 
"infinite P-sink" in the sense that P release by the soil is clearly the rate-limiting step 
(Sibbesen, 1978; van der Zee et al., 1987; Yli-Halla, 1990). For anion-exchange resins 
used at low resin:soil ratios, this relationship cannot be assumed (Barrow and Shaw, 
1977; Pierzynski, 1991) and is not necessary for the assessment of bioavailable P. 



39 



Methods for P Analysis, G.M. Pierzynski (ed) 

Iron-oxide-Impregnated Paper 

Another P sink that has received attention is Fe-oxide impregnated filter paper, which 
has successfully estimated plant available P in a wide range of soils and management 
systems (Menon et al, 1989; 1990, Sharpley, 1991). Also, Sharpley (1993) observed that 
the Fe-oxide strip P content of runoff was closely related to the growth of several algal 
species incubated for 29-d with runoff as the sole source of P. As the resin membranes 
and Fe-oxide strips act as a P sink, they simulate P removal from soil or sediment-water 
samples by plant roots and algae. Thus, they have a stronger theoretical justification for 
use over chemical extractants to estimate bioavailable P. These methods have potential 
use as environmental soil P tests to identify soils liable to enrich runoff with sufficient P 
to accelerate eutrophication. The Fe-oxide impregnanted filter paper procedure was 
described in the section by Chardon (2000) in this bulletin and will not be described 
further here. 

Anion-exchange Resins 

The use of anion-exchange resins is the most common P-sink approach for assessing 
available inorganic P in soils. The procedure typically involves the use of chloride- 
saturated resin at a 1:1 resin-to-soil ratio in 10 to 100 mL of water or weak electrolyte for 
16 to 24 h (Amer et al., 1955; Olsen and Sommers, 1982). Correlations between plant 
response and resin-extractable P are comparable or superior to correlations with chemical 
extraction methods (Fixen and Grove, 1990). 

Ion-exchange Resin-Impregnated Membranes 

A similar approach using ion-exchange resin impregnated membranes has been 
investigated by several researchers (Abrams and Jarrell, 1992; Qian et al., 1992: Saggar 
et al., 1992). Impregnation of the resin onto a plastic membrane facilitates separation of 
the resin beads from the soil and may eliminate the soil grinding step. Also, an extraction 
time as short as 15 min can be used without reducing the accuracy of predicted P 
availability for a wide range of soils (Qian et al., 1992). In pot studies, the resin 
membranes have provided a better index of P availability than conventional chemical 
extraction methods for canola (Qian et al., 1992) and ryegrass (Saggar et al., 1992). It is 
likely that the utility of the resin membranes will make the use of loose resin obsolete. 

Ion exchange membranes have the potential to estimate P availability in aquatic as 
well as soil environments. Edwards et al. (1993) used ion exchange membranes to obtain 
in-situ estimates of the chemical composition of river water for two Scottish watersheds. 
It was suggested that direct multi-element analysis by X-ray fluorescence of ions retained 
on the membranes removes the need for sample storage or filtration, both of which can be 
sources of potential contamination and error. Thus, the membranes can provide useful 
information in addition to that obtained by conventional sampling (Edwards et al., 1993). 

Soil Sampling: 

Soil sampling protocol for environmental concerns should be re-evaluated since the 
primary mechanism for P transport from most agricultural soils is by surface runoff and 
erosion. Although most samples submitted to soil testing laboratories are obtained from 
to 20 cm, the zone of interaction of runoff waters with most soils is normally less than 5 
cm. Consequently, environmental soil sampling should reflect this shallower depth of 



40 



Methods for P Analysis, G.M. Pierzynski (ed) 

soil influencing runoff P. Hence, environmental soil samples should, in general, be taken 
from no deeper than 5 cm. This protocol is compatible with sampling of no-till fields, 
currently recommended by extension specialists in several states, where the traditional 0- 
to 20-cm depth is split into two or three increments. Thus, on soils identified as 
vulnerable to P loss in runoff, the surface increment could be analyzed for environmental 
interpretation and all increments integrated for agronomic interpretations. 

Equipment: 

The following equipment is needed to conduct BAP extraction of soil and analysis for 
P: 

1. Resin membrane, anion exchange. 

2. End-over-end shaker - used to equilibrate sample and sink 

3. Volumetric flasks - usually 25 or 50 mL volume 

4. Pipets to aliquot samples and color reagents 

5. Spectrophotometer to determine P concentration in the color developed reagent 
with sample. 

Reagents: 

Resin membranes 

1. Hydrochloric acid to extract P from the membranes - 1.0 M HC1 (166 mL 
concentrated HC1 in 2 L) 
Murphy and Riley Molybdenum Blue Color Reagent 

1. Murphy and Riley Reagent A: 

a. 1. Mix 1500 mL H 2 and 125 mL H2SO4 and allow to cool down before 
adding molybdate and tartrate 

b. Add 10.66 g ammonium molybdate 

c. Add 50 mL antimony potassium tartrate 

d. Make the solution up to 2 L 

e. Store in refrigerator 

2. Murphy and Riley Reagent B: 

a. Dissolve 42 g ascorbic acid in 1 L 

b. Store in refrigerator 

3. Murphy and Riley Reagent 

The color development reagent is made up by mixing nine parts of reagent A and 
1 part of reagent B in a measuring cylinder. Each sample in a 25 mL volumetric 
flask requires 5 mL of this reagent. As it takes time to make up the Murphy and 
Riley reagent and some of the reagents are expensive (e.g., ammonium 
molybdate), only make up what is needed for the day. Also, solutions A and B, 
once mixed, will not keep for more than a day. For example, if you have 20 
samples to run this will require at least 100 mL of color reagent plus standards 
and some for reruns. Thus, 250 mL of color reagent should be mixed, and this 
will require 225 mL of reagent A and 25 mL of reagent B. 

4. Neutralizing Reagents : 

a. p-nitrophenol indicator (pnp - yellow): mix 1.5 g p-nitrophenol in 500 mL of 
deionized distilled water on a magnetic stirrer until dissolved. Filter the 
solution to remove any undissolved residue. 



41 



Methods for P Analysis, G.M. Pierzynski (ed) 

b. 4 M NaOH: 160 g NaOH in 1 L 

c. 0.1MH 2 SO 4 : 11.1 mL cone. H2SO4 in 2 L 
5. Solution Neutralizing 

a. Add one drop of pnp indicator to an appropriate aliquot of the filtered solution 
on which P is to be measured in a volumetric flask. 

b. Add 4 M NaOH to solution drop-wise until solution just turns yellow. 

c. Add 0.1MH; SO4 drop-wise until solution just turns back to clear, the 
solution is now neutral and the Murphy and Riley reagent can be added. 

Resin Strip Procedure: 

1 . Anion exchange resin sheets are cut into 2x2 cm squares and are stored in 
propylene glycol. Wash the resin squares in distilled water to remove all the 
propylene glycol. If not already saturated with an anion, saturation with CI , 
HCO3" or acetate may be necessary. They are now ready for use. 

2. Phosphorus is extracted from soil or sediment by shaking a 1-g sample and one 
resin membrane square in 40 mL of deionized distilled water end-over-end for 16 
hours at 25° C. 

3. Remove the resin membrane square and wash thoroughly with distilled water until 
all soil particles are removed. 

4. The BAP content of runoff can also determined by shaking 50 mL of an unfiltered 
runoff sample with one resin membrane square for 16 hours. Smaller runoff 
sample volumes should be used if P concentrations are expected to be high (>1 or 
2 mg/L) and made up to 50 mL with distilled water. 

5. Phosphorus retained on the resin membrane square is removed by shaking the 
square end-over-end with 40 mL of 1 M HC1 for 4 hours. Remove square and 
rinse with distilled water. Retain the HC1 desorption solution for analysis. Repeat 
this step. Do not mix the first and second desorption solutions. 

6. Measure the P concentration of the two solutions separately. The total amount of 
P desorbed from the resin membrane square is the sum of the amounts in the two 
solutions. 



Calculations: 

Resin extractable P (mg P/kg) = 

[Concentration of P in 1 MHC1, mg/L] x [0.04 L 4- 0.001 kg] 

Resin BAP in runoff (mg P/L) = [concentration of P in 1 M HC 1 , mg/L] x [0.04L ■ 
volume of runoff, L] 



References: 

Abrams, M.M., and W.M. Jarrell. 1992. Bioavailability index for phosphorus using 
nonexchange resin impregnated membranes. Soil Sci. Soc. Am. J. 56:1532-1537. 

Amer, F., D.R. Bouldin, C.A. Black, and F.R. Duke. 1955. Characterization of soil 
phosphorus by anion exchange resin and adsorption by P-32 equilibration. Plant Soil 
6:391-408. 



42 



Methods for P Analysis, G.M. Pierzynski (ed) 

Barrow, N.J., and T.C. Shaw. 1977. Factors affecting the amount of phosphate extracted 

from soil by anion exchange resin. Geoderma 18:309-323. 
Butkus, S.R., E.B. Welch, R.R. Horner, and D.E. Spyridakis. 1988. Lake response 

modeling using biologically available phosphorus. J. Water Pollut. Cont. Fed. 

60:1663-1669. 
Chardon, W.J. 2000. Phosphorus extraction with iron oxide-impregnated filter paper (P; 

test). In G.M. Pierzynski (ed.), Methods for Phosphorus Analysis for Soils, 

Sediments, Residuals, and Waters. Southern Cooperative Series Bulletin No. 396, p. 

27-30. 
Dorich, R.A., D.W. Nelson, and L.E. Sommers. 1980. Algal availability of sediment 

phosphorus in drainage water of the Black Creek watershed. J. Environ. Qual. 9:557- 

563. 
Edwards, T., B. Ferrier, and R. Harriman. 1993. Preliminary investigation on the use of 

ion-exchange resins for monitoring river water composition. Sci. of Total Environ. 

135:27-36. 
Fixen, P.E., and J.H. Grove. 1990. Testing soils for phosphorus, p. 141-180. InR.L. 

Westerman (ed.) Soil testing and plant analysis. 3rd ed. SSSA Book Ser. 3. SSSA, 

Madison, WI. 
Hope, G.D., and J.K. Syers. 1976. Effects of solution to soil ratio on phosphate sorption 

by soils. J. Soil Sci. 27:301-306. 
Huettl, P.J., R.C. Wendt, and R.B. Corey. 1979. Prediction of algal available 

phosphorus in runoff suspension. J. Environ. Qual. 4:541-548. 
Logan, T.J., T.O. Oloya, and S.M. Yaksich. 1979. Phosphate characteristics and 

bioavailability of suspended sediments from streams draining into Lake Erie. J. Great 

Lakes. Res. 5:112-123. 
Menon, R.G., L.L. Hammond, and HA. Sissingh. 1989. Determination of plant- 
available phosphorus by the iron hydroxide-impregnated filter paper (Pi) soil test. 

Soil Sci. Soc. Am. J. 52:110-115. 
Menon, R.G., S.H. Chien, L.L. Hammond, and B.R. Arora. 1990. Sorption of 

phosphorus by iron oxide-impregnated filter paper (Pi soil test) embedded in soils. 

Plant Soil 126:287-294. 
Miller, W.E., J.C. Greene, and T. Shiroyarna. 1978. The Selenastrum capricornutum 

Printz algal assay bottle test and data interpretation protocol. U.S. EPA, Tech. Rep. 

EPA-600/9-78-018. USEPA, Covallis, OR. 
Olsen, S.R., and L.E. Sommers. 1982. Phosphorus, p. 403-429. In A.L. Page et al. 

(eds.), Methods of soil analysis. Agronomy No. 9, Part 2, 2nd ed., Am. Soc. of 

Agronomy, Madison, WI. 
Pierzynski, G.M. 1991. The chemistry and mineralogy of phosphorus in excessively 

fertilized soils. Critical Reviews in Environ. Control 21:265-295. 
Porcella, D.B., J.S. Kumazar, and E.J. Middlebrooks. 1970. Biological effects on 

sediment-water nutrient interchange. J. Sanit. Eng. Div., Proc. Am. Soc. Civil Eng. 

96:911-926. 
Qian, P., J.J. Schoenau, and W.Z. Huang. 1992. Use of ion exchange membranes in 

routine soil testing. Commun. Soil Sci. Plant Anal. 23:1791-1804. 



43 



Methods for P Analysis, G.M. Pierzynski (ed) 

Saggar, S., MJ. Hedley, R.E. White, P.E.H. Gregg, K.W. Perrot, and I.S. Conforth. 

1992. Development and evaluation of an improved soil test for phosphorus. 2. 

Comparison of the Olsen and mixed cation-anion exchange resin tests for predicting 

the yield of ryegrass growth in pots. Fert. Res. 33: 135-144. 
Sharpley, A.N. 1991. Soil phosphorus extracted by iron- aluminum-oxide-impregnated 

filter paper. Soil Sci. Soc. Am. J. 55:1038-1041. 
Sharpley, A.N. 1993. An innovative approach to estimate bioavailable phosphorus in 

agricultural runoff using iron oxide-impregnated paper. J. Environ. Qual. 22:597- 

601. 
Sharpley, A.N., W.W. Troeger, and S.J. Smith. 1991. The measurement of bioavailable 

phosphorus in agricultural runoff. J. Environ. Qual. 20:235-238. 
Sharpley, A.N., L.R. Ahuja, M. Yamamoto, and R.G. Menzel. 1981. The kinetics of 

phosphorus desorption from soil. Soil Sci. Soc. Am. J. 45:493-496. 
Sibbesen, E. 1978. An investigation of the anion-exchange resin method for soil 

phosphate extraction. Plant Soil 50:305-321. 
Sonzogni, W.C., S.C. Chapra, D.E. Armstrong, and T.J. Logan. 1982. Bioavailability of 

phosphorus inputs to lakes. J. Environ. Qual. 11:555-563. 
van der Zee, S.E.A.T.M., L.G.J. Fokkink, and W.H. van Riemsdijk. 1987. A new 

technique for assessment of reversibly adsorbed phosphate. Soil Sci. Soc. Am. J. 

51:599-604. 
Yli-Halla, M. 1990. Comparison of a bioassay and three chemical methods for 

determination of plant-available P in cultivated soils of Finland. J. Agric. Sci. Finl. 

62:213-319. 



44 



Methods for P Analysis, G.M. Pierzynski (ed) 

Total Phosphorous in Soil 

M.R. Bender and C.W. Wood, Auburn University 



Introduction: 

There have been many methods developed to extract and analyze total phosphorus (P) 
in soil (Bray and Kurtz, 1945; Muir, 1952; Jackson, 1958; Syers et al., 1968; Sommers 
and Nelson, 1972; Dick and Tabatabai, 1977; Olsen and Sommers, 1982; Bowman, 
1988). Two of the more commonly used and most recognizable methods of P extraction 
are sodium carbonate (Na2C03) fusion and acid digestion. Of these methods, Na2CC>3 
fusion is thought to give more reliable results (Syers et al., 1967; Syers et al., 1968; 
Sherrell and Saunders, 1966; Sommers and Nelson, 1972). Underestimation of total P by 
acid digestion is thought to be due to inability of these methods to extract P from apatite 
inclusions (Syers et al., 1967). The ability of an acid digestion to extract P from 
inclusions depends upon the acid or combination of acids used. Syers et al. (1967) 
showed that the effectiveness of extraction generally followed the order: fusion > HF 
digestion > HCIO4 digestion >iVH 2 SO4 > ignition. 

In recent years, more rapid methods for determining total P in soils have been 
developed (Sommers and Nelson, 1972; Dick and Tabatabai, 1977; Bowman, 1988). 
Methods developed by Sommers and Nelson (1972) and Bowman (1988) are variations 
of standard HCIO4 digestion methods. These methods were shown to give a similar 
degree of underestimation of total P as standard HCIO4 digestion methods. Dick and 
Tabatabai (1977) proposed an alkaline oxidation method using sodium hypobromite 
(NaOBr). This method was shown to give results 1% higher than those found by HCIO4 
digestion. However, the method still underestimated total P by 4% when compared to 
results from Na2CC>3 fusion. 

The methods discussed here are very silimar to Na2CC>3 fusion and HCIO4 digestion as 
described by Olsen and Sommers (1982) in Methods of Soil Analysis - Part 2, and the 
alkaline oxidation method developed by Dick and Tabatabai (1977). 

Fusion Method (Olsen and Sommers (1982)): 

Reagents 

1. Anhydrous sodium carbonate (Na2C03) 

2. 4.5MH 2 S0 4 

3. IMH2SO4 

4. Ammonium paramolybdate [(NH 4 ) 6 Mo7024 'H 2 0]. Prepare by dissolving 9.6 g of 
(NH 4 ) 6 Mo7 024 ' 4H 2 in distilled water under heat. After solution has cooled, 
dilute solution volume to 1 L with distilled water. 

5. 2MH2SO4 

6. Ascorbic acid. Prepare by dissolving 10 g of ascorbic acid in 80 mL of distilled 
water, and dilute solution volume to 100 mL with distilled water. Store reagent at 

o 

2 C. Make fresh solution when noticeable color develops. 

7. Potassium antimony tartrate (KSbOC4H 4 06). Prepare by dissolving 0.667 g of 
KSbO-C 4 H 4 6 in 250 mL of distilled water. 

8. Mixed reagent. Mix 1:1 ratio of ascorbic acid and antimony reagents prior to use. 
Prepare a fresh solution as required. 



45 



Methods for P Analysis, G.M. Pierzynski (ed) 



Procedure 

Place a mixture of 1.0 g of finely ground (100 mesh), air-dried soil and 4-5 g of 
Na2CC>3 in a Pt crucible. For soils high in Fe, use 0.5 g of soil. Place 1 g of Na2CC>3 on 
top of the mixture. Drive off moisture from mixture by gently heating with a Meeker 
burner. Place a lid on the crucible so that approximately one fifth of the crucible remains 
open. Apply heat with a low flame for 10 min so the mass fuses gently. Adjust heat of 
Meeker burner to full, and heat mass for 15 to 20 min. To provide an oxidizing 
environment for this step, lift the lid of the crucible periodically. Do not allow the 
reduced portion of the flame to come in contact with the crucible. Remove crucible from 
flame. Rotate crucible as it cools so to deposit the melt thinly onto the walls of the 
crucible. After the crucible has cooled, gently roll it between your hands to facilitate the 
removal of the melt. Remove the melt with 30 mL of 4.5 M H2SO4, using care to avoid 
loss by effervescence. Place crucible and lid in a beaker containing 25 mL of 1 M H2SO4, 
and heat contents to a boil. Transfer the solution from the beaker and the solution from 
the melt to a 250 mL volumetric flask. Dilute the solution to volume using distilled water. 
Allow sediment to settle. Remove an aliquot of clear supernatant solution for total P 
analysis by the ascorbic acid method. 

To analyze for total P, transfer aliquots (2 mL) into 50 mL volumetric flasks (for 
samples containing <150 mg of P). With 1 M Na2CC>3 , adjust pH of the aliquot to 5 using 
p-nitrophenol indicator. Add 5 mL of 2 M H 2 SO4 and 5 mL of ammonium 
paramolybdate reagent and mix. Add 4 mL of the mixed reagent and mix contents of the 
flask. Bring to 50 mL volume with distilled water and mix thoroughly. Reduction is 
completed and maximum color intensity develops in 10 min, and color is stable for 24 
hours. The absorption maximum of the blue color formed in the presence of Sb is at 890 
nm (Harwood et al., 1969) 

Comments 

The method for color development was described by Harwood et al. (1969) and is a 
variation of the method proposed by Murphy and Riley (1962). By increasing amount of 
antimony added, Harwood et al. (1969) found that the range of the calibration curve 
could be extended. This modification of the Murphy and Riley (1962) method was found 
to increase the upper limit of the calibration curve from 50 mg P/50ml sample to 150 mg 
P/50ml sample. 

It should be noted that presence of arsenic in the form of ASO4 in soil samples 
gives the same blue color as phosphate. To eliminate this problem, ASO4 can be reduced 
to ASO3 using a NaHSC>3 solution as described in the following digestion method (Olsen 
and Sommers, 1982). 

Calculations 

Total P, mg/kg = 

[Concentration of P in initial 250 mL dilution, mg/L] x [0.25 L 4- mass of soil, kg] 



46 



Methods for P Analysis, G.M. Pierzynski (ed) 

Digestion Method (Olsen and Sommers (1982)): 

Reagents 

1. 60% Perchloric acid (HC10 4 ) 

2. Ammonium paramolybdate-vanadate. Prepare by dissolving 25 g of (NH 4 ) 6 Mo7024 
' 4H 2 O in 400 mL of distilled water, and by dissolving ammonium metavanadate 
(NH4VO3) in 300 mL of boiling distilled water. Cool vanadate solution, and add 
250 mL of cone. HNO3 . Cool NH4VO3-HNO3 solution to room temperature before 
adding (NH 4 )6Mo7 024'4H 2 solution. Dilute the mixed solution to 1 L with 
distilled water. 

3. Standard phosphate solution. Prepare by dissolving 0.4393 g of oven-dried 
potassium dihydrogen phosphate (KH2PO4) in distilled water. Dilute solution to 1 
L with distilled water. Standard solution contains 100 mg P/L. 

4. Sodium hydrogen sulfite (NaHSC^). Prepare by dissolving 5.2 g of reagent grade 
NaHS0 3 in 100 mL of 0.5 M H 2 S0 4 . Prepare reagent weekly. 

Procedure 

In a 250 mL volumetric or Erlenmeyer flask, mix 2.0 g of finely ground soil (<0.5 
mm) with 30 mL of 60% HCIO4 . Digest the soil and acid mixture at a few degrees below 
the boiling point on a hot plate in a perchloric hood until the dark color from organic 
matter disappears. Continue to heat at the boiling temperature for 20 min longer. Heavy 
white fumes will appear, and the insoluble material will become like white sand. If any 
black particles stick to the side of the flask, add 1 or 2 mL of HCIO4 to wash down the 
particles. If the sample is high in organic matter it may be necessary to add 20 mL of 
HNO3 and heat to oxidize organic matter before adding HCIO4 . Total digestion time is 
approximately 40 min. Cool the mixture before bringing the volume up to 250 mL with 
distilled water. Mix the contents of the flask, and then allow sediment to settle. 

To analyze for total P, transfer aliquots into 50 mL volumetric flasks (for samples 
containing between 0.05 to 1.0 mg of P). Add 10 mL of the ammonium paramolybdate- 
vanadate reagent, and bring the volume of the flask up to 50 mL using distilled water. 
The optical density of the sample can be measured after 10 min at wavelengths between 
400 to 490 nm. The optical density of a reagent blank should be subtracted from the 
optical density readings of the samples. 

To reduce As(V 3 to As(V 3 , add 5 mL of NaHS03 solution to the aliquot. Then 
partially immerse the 50 mL volumetric flasks in a water bath, and digest the solution for 
30 min (20 min after temperature reaches 95°C). An alternative procedure is to allow the 
solution to stand for 4 hours at room temperature. 

Calculations 

Total P, mg/kg = 

[Concentration of P in initial 250 mL dilution, mg/L] x [0.25 4- mass of soil, kg] 



47 



Methods for P Analysis, G.M. Pierzynski (ed) 

Alkaline Oxidation Method (Dick and Tabatabai (1977)): 

Reagents 

1. Sodium hypobromite solution (NaOBr-NaOH). Prepare by slowly adding 3 mL of 
bromine (0.5 mL/min) to 100 mL of 2 M NaOH under constant stirring. Prepare 
reagent immediately prior to use. 

2. 90 % formic acid 

3. 2.5MH 2 S0 4 

4. Ammonium molybdate -Antimony potassium tartrate solution. Prepare by 
dissolving 12 g of ammonium molybdate in 250 mL of distilled water, and 
dissolving 0.2908 g of antimony potassium tartrate in 100 mL of distilled water. 
Add both solutions to 1 L of 2.5 M sulfuric acid, and dilute volume to 2 L with 
distilled water. Store reagent in a cool place, in a dark Pyrex glass bottle. 

5. Ascorbic acid. Prepare by dissolving 1.056 g of ascorbic acid in 200 mL of 
ammonium molybdate - antimony reagent. Prepare reagent daily. 

6. Standard phosphate solution. Prepare by dissolving 0.2195 g of potassium 
dihydrogen phosphate (KH2PO4) in distilled water. Dilute solution to 1L with 
distilled water. Standard solution contains 50 mg P/L. 

Procedure 

Place a 100 to 200 mg sample of finely ground, air-dried soil in a 50 mL boiling flask. 
Add 3 mL of sodium hypobromite solution to the flask, and swirl flask for a few seconds 
to mix contents. Allow flask to stand for 5 min. Swirl flask again and place it in a sand 
bath adjusted to 260 to 280°C. The sand bath should be situated in a hood. Heat flask 
until contents evaporate to dryness. Evaporation time is 10 to 15 min. After evaporation, 
continue to heat for an additional 30 min. Remove flask from sand bath, and allow it to 
cool for 5 min. Then add 4 mL of distilled water and 1 mL of formic acid. Mix contents 
before adding 25 mL of 0.5 M H 2 SO4 . Stopper flask and mix contents. Transfer mixture 
to a 50 mL plastic centrifuge tube and centrifuge sample at 12,000 rpm for 1 min. 

To analyze for total P, transfer aliquots of 1 to 2 mL into 25 mL volumetric flasks. 
Add 4 mL of ascorbic acid reagent, and bring solution up to volume with distilled water. 
Stopper flask and mix solution. Allow solution to stand for 30 min for color development. 
Optical density of sample should be measured at a wavelength of 720 nm. 

Comments 

This method does not require neutralization of the 1 to 2 mL of aliquot, however, 
longer time (30 min) is needed for full color development. 

The sodium hypobromite (NaOBr-NaOH) reagent should be prepared just prior to use. 
The reagent should be made in a fume hood. Formic acid added after the hypobromite 
treatment will destroy any residual hypobromite remaining after oxidation of the sample. 

Calculations 

Total P, mg/kg = 

[Concentration of P in initial formic acid/H 2 S04 solution, mg/L] x [0.03 L 4- mass of 
soil, kg] 



48 



Methods for P Analysis, G.M. Pierzynski (ed) 

References: 

Bowman, R.A. 1988. A rapid method to determine total phosphorus in soils. Soil Sci. 

Soc. Am. J. 52:1301-1304. 
Bray, R.H., and L.T. Kurtz. 1945. Determination of total, organic, and available forms of 

phosphorus is soils. Soil Sci. 59:39-45. 
Dick, W.A., and M.A. Tabatabai. 1977. An alkaline oxidation method for determination 

of total phosphorus in soils. Soil Sci. Soc. Am. J. 41:511-514. 
Harwood, J.E., R.A. van Steenderen, and A.L. Kuhn. 1969. A rapid method for 

orthophosphate analysis at high concentrations in water. Water Res. 3:417-423. 
Jackson, M.L. 1958. Soil chemical analysis. Prentice-Hall, Inc., Englewood Cliffs, N.J. 
Muir, J.W. 1952. The determination of total phosphorus in soil. Analyst 77:313-317. 
Murphy, J., and J.P. Riley. 1962. A modified single solution method for determination of 

phosphate in natural waters. Anal. Chim. Acta 27:31-36. 
Olsen, S.R., and L.E. Sommers. 1982. Phosphorus, pp. 403-430. In: A.L. Page. R.H. 

Miller, and D.R. Keeney (eds.), Methods of Soil Analysis. 2nd ed. Agronomy Series 

No. 9, Part 2. Soil Science Society of America, Inc., Madison, WI. 
Sherrell, C.G., and W.M.H. Saunders. 1966. An evaluation of methods for the 

determination of total phosphorus in soils. N.Z.J. Agric. Res. 9:972-979. 
Sommers, L.E., and D.W. Nelson. 1972. Determination of total phosphorus in soils: a 

rapid perchloric acid digestion procedure. Soil Sci. Soc. Amer. Proc. 36:902-904. 
Syers, J.K., J.D.H. Williams, A.S. Campbel, and T.W. Walker. 1967. The significance of 

apatite inclusions in soil phosphorous studies. Soil Sci. Soc. Amer. Proc. 31:752-756. 
Syers, J. K., J.D.H. Williams, and T.W. Walker. 1968. The determination of total 

phosphorus in soils and parent materials. N.Z.J. Agric. Res. 11:757-762. 



49 



Methods for P Analysis, G.M. Pierzynski (ed) 



Phosphorus Fractionation 

Hailin Zhang, Oklahoma State University 
John L. Kovar, USDA/ARS, Ames, IA 



Introduction: 

The chemistry of phosphorus (P) in soils is complicated. Inorganic P can react with 
Ca, Fe and Al to form discrete phosphates, and organic P can be in different forms with 
varying resistance to microbial degradation. To investigate the forms of inorganic P (P ; ) 
and transformations of applied P fertilizers, the fractionation procedure of Chang and 
Jackson (1957) has been widely used. Subsequent studies indicated that various 
extractants were not as specific as first envisioned. For example, retention of P by CaF2 
formed from CaC03 during ammonium flouride (NH4F) extraction affects results when 
the Chang and Jackson method is used with calcareous soils and sediments. Since its 
development, modifications made by Williams et al. (1967), Smillie and Syers (1972), 
Peterson and Corey (1966), and Fife (1962) have improved extractability and allowed for 
use with calcareous soils. The original fractionation procedures and the most important 
modifications were summarized by Kuo (1996). The Pj fractionation in this paper is 
primarily based on the Kuo (1996) fractionation scheme. 

Soil organic P (P ) consists of inositol phosphates, phospholipids, nucleic acids, 
phosphoproteins, and various sugar phosphates, as well as a significant number of 
compounds that have not been identified. Organic P tied up in microbial biomass 
consists of nucleic acids, inositol phosphates, and polyphosphates. Microbial biomass P 
usually represents a small fraction of the total P in soil, and rapidly turns over to supply 
inorganic P to plant roots (Tate, 1984). Quantification of the various known P 
compounds in soil has been described in several studies (Anderson, 1967; Halstead and 
Anderson, 1970; Stott and Tabatabai, 1985) and is advocated by Kuo (1996) as a means 
of fractionating soil P . An alternative method for characterizing soil P fractions 
involves the use of acid and alkaline extractants that separate the various fractions based 
on the type and strength of P physicochemical interactions with other soil components 
(Bowman and Cole, 1978; Hedley et al., 1982; Cross and Schlesinger, 1995). The most 
common extractants are 0.5 M sodium bicarbonate (NaHCCh) and various concentrations 
of hydrochloric acid (HC1) and sodium hydroxide (NaOH). The fractionation scheme 
involves a sequence of extractions that separates soil P into labile, moderately labile, and 
nonlabile fractions. In recent years, this scheme has been widely used to evaluate P 
turnover in diverse soils under varying management (Hedley et al., 1982; Sharpley and 
Smith, 1985; Ivanoff et al., 1998). The qualitative and quantitative information provided 
by the fractionation data is useful for agronomic and water quality issues. 

Fractionation of Inorganic Phosphorus: 

Principles 

The fractionation procedures are based on the differential solubilities of the various 
inorganic P forms in various extracts. Ammonium chloride (NH4CI) is used first to 
remove soluble and loosely bound P, followed by separating Al-P from Fe-P with 
(NH4F), then removing Fe-P with NaOH. The reductant- soluble P is removed with CDB 



50 



Methods for P Analysis, G.M. Pierzynski (ed) 

(sodium citrate- sodium dithionite- sodium bicarbonate) extraction. The Ca-P is extracted 
with sulfuric acid ( H2SO4 ) or HC1 since Ca-P is insoluble in CDB. Since NH 4 F reacts 
with CaC03 to form CaF 2 in calcareous soils, which will precipitate soluble P and reduce 
the effectiveness of NH4F to extract P, the NH4F extraction is omitted for calcareous 
soils. 

Equipment 

1. Shaker 

2. Centrifuge and 100-mL centrifuge tubes 

3. Hot water bath 

4. Spectrophotometer 

Reagents 

1. \ M ammonium chloride (NH 4 C1). Dissolve 53.3 g of NH 4 C1 in 1 L deionized 
water 

2. 0.5 M ammonium fluoride (NH 4 F) pH 8.2. Dissolve 18.5 g of NH 4 F in 1 L 
deionized water and adjust pH to 8.2 with 4 M NH 4 OH. 

3. 2 M and 0.1 M sodium hydroxide (NaOH). Dissolve 80 g and 4.0 g respectively of 
NaOH in 1 L deionized water. 

4. 0. 1 M NaOH + 1 M NaCl. Dissolve 4.0 g of NaOH and 58.5 g of NaCl in 1 L 
deionized water. 

5. Saturated NaCl. Add 400 g of NaCl to 1 L deionized water. 

6. 0.25 M sulfuric acid. Dilute 14 mL of concentrated H 2 SO4 to 1 L with deionized 
water. 

7. 2 M hydrochloric acid. Dilute 168 mL of concentrated HC1 to 1 L with deionized 
water. 

8. 0.3 M sodium citrate. Dissolve 88.2 g of Na3C6H 5 07'2H 2 in 1 L deionized water. 

9. \M sodium bicarbonate. Dissolve 84 g of NaHC03 in 1L deionized water. 

10. 0.8 M boric acid. Dissolve 50 g of H3BO3 in 1 L deionized water. 

11. Sodium dithionite reagent grade. 

12. 0.25% p-nitrophenol. Dissolve 0.25 g of p-nitrophenol in 100 mL of deionized 
water. 

Procedures for Noncalcareous Soils (flow chart in Fig. 1) 

1 . Add 1 .0 g (<2 mm) of soil and 50 mL of \M NH 4 C1 to a 100 mL centrifuge tube 
and shake for 30 min to extract the soluble and loosely bound P. Centrifuge and 
decant the supernatant into a 50-mL volumetric flask and bring to volume with 
deionized water (extract A). 

2. Add 50 mL of 0.5 M NH 4 F (pH 8.2) to the residue and shake the suspension for 1 
h to extract aluminum phosphates. Centrifuge and decant the supernatant into a 
100-mL volumetric flask (extract B). 

3. Wash the soil sample twice with 25-mL portions of saturated NaCl and centrifuge. 
Combine the washings with extract B and bring to volume. Add 50 mL of 0.1 M 
NaOH to the soil residues and shake for 17 h to extract iron phosphate. Centrifuge 
and decant the supernatant solution into a 100-mL volumetric flask (Extract C). 
Wash the soil twice with 25-mL portions of saturated NaCl and centrifuge. 
Combine the washings with extract C and bring to volume. 



51 



Methods for P Analysis, G.M. Pierzynski (ed) 



Noncalcareous Soils 



Calcareous Soils 



1.0 g of soil in 100 mL 
centrifuge tube 



50 mL 1 M NH 4 C1, shake 
30 min., centrifuge 



50 mL 0.5 M NH 4 F, shake 

1 hr., centrifuge, wash with 

saturated NaCl 



Soluble and loosely 



bound P 



Al-P 











50 mL 0.1 M NaOH, shake 
17 hrs., centrifuge and wash 


Fe-P 











40 mL 0.3M Na 3 C 3 H 6 0,, 5 

mL 1 M NaHCOj, 1.0 g 

Na,S,0 4 , heat, stir, heat, 

centrifuge and wash 



Reductant soluble 

"* P 



50 mL 0.25 M H,S0 4 , 

shake 1 hr., centrifuge and 

wash 



Ca-P 



1.0 g of soil in 100 mL 
centrifuge tube 



50 mL 0.1 M NaOH + 1 M 
NaCl, shake 17 hrs., 
centrifuge and wash 



40 mL 0.3M Na 3 CjH 6 7 , 5 

mL 1 M NaHCOj, 1.0 g 

Na,S,0 4 , heat, stir, heat, 

centrifuge and wash 





^ 


' 




50 mL 0.5 M HC1, shake 1 
hr., centrifuge and wash 













Figure 1. Sequential fractionation scheme for inorganic P. 



52 



Methods for P Analysis, G.M. Pierzynski (ed) 



4. Add 40 mL of 0.3 M Na 3 C 6 H 5 7 and 5 mL of 1 M NaHC0 3 to the residue and heat 
the suspension in a water bath at 85 °C. 

5. Add 1.0 g of Na2S2C>4 (sodium dithionate) and stir rapidly to extract reductant- 
soluble P. Continue to heat for 15 min and then centrifuge. Decant the supernatant 
solution into a 100-mL volumetric flask (extract D). Wash the soil twice with 25- 
mL portions of saturated NaCl and centrifuge. Combine the washings with extract 
D, and dilute D to volume. Expose extract D to air to oxidize NaiSiCv 

6. Add 50 mL of 0.25 M H 2 SO4 to the soil residue and shake for 1 h. Centrifuge the 
suspension for 10 min and decant the supernatant into a 100-mL volumetric flask 
(extract E). Wash the soil twice with 25-mL portions of saturated NaCl, and 
centrifuge. Combine the washings with the extract E and dilute to volume. 

7. Transfer an aliquot containing 2 to 40 (ig P from each of extracts A, B, C, D, and E 
to separate 50-mL volumetric flasks. Add some deionized water and five drops of 
p-nitrophenol indicator to the volumetric flasks containing extracts C and E, and 
adjust the pH with 2 M HC1 or 2 M NaOH until the indicator color just changes. 
Add 15 mL 0.8 M H3BO3 to the volumetric flask containing extract B. Phosphorus 
concentrations in the various solutions can be determined using the ascorbic acid 
method (Murphey and Riley, 1962). Prepare P standards that contain the same 
volume of extracting solution as in the extracts. 

Calculations 

The amount of P in each fraction is calculated using the following equation: 

P concentration in given fraction (mg/kg) = 
[Cone, of P (mg/L)] x [Volume of extractant (L) 4- mass of soil (kg)] 

Procedures for Calcareous Soils (flow chart in Fig. 1) 

1. Add 1.0 g (<2 mm) of soil and 50 mL of 0.1 M NaOH + 1 M NaCl, shake for 17 
h. Centrifuge and decant the supernatant solution into a 100-mL volumetric flask 
(extract A). Wash the soil twice with 25-mL portions of saturated NaCl and 
centrifuge. Combine the washings with extract A and bring to volume. 

2. Add 40 mL of 0.3 M Na3C 6 H 5 07 and 5 mL of 1 M NaHC0 3 to the residue and heat 
the suspension in a water bath at 85°C. Add 1.0 g of Na2S2 04 (sodium dithionate) 
and stir rapidly. Continue to heat for 15 min and centrifuge. Decant the supernatant 
solution into a 100-mL volumetric flask (extract B). Wash the soil twice with 25- 
mL portions of saturated NaCl and centrifuge. Combine the washings with extract 
B, and dilute B to volume. Expose extract B to air to oxidize Na2S2C>4. 

3. Add 50 mL of 0.5 M HC1 to the soil residue and shake for 1 h. Centrifuge the 
suspension, and decant the supernatant into a 100-mL volumetric flask (Extract C). 
Wash the soil twice with 25-mL portions of saturated NaCl, and centrifuge. 
Combine the washings with the extract C and dilute to volume. 

4. Transfer an aliquot containing 2 to 40 (ig P from each of extracts A, B, and C to 
separate 50-mL volumetric flasks. Add some deionized water and five drops of p- 
nitrophenol indicator to each of the volumetric flasks containing extracts A and C 
and adjust the pH with 2 M HC1 or 2 M NaOH until the indicator color just 



53 



Methods for P Analysis, G.M. Pierzynski (ed) 

changes from yellow to colorless for extracts A and C. P concentrations of various 
fractions can be determined using the ascorbic acid method (Murphey and Riley, 
1962). Prepare P standards that contain the same volume of extracting solution as 
in the extracts. 

Calculations 

The amount of P in each fraction can be calculated using the following equation: 

P concentration in given fraction (mg/kg) = 

[Cone, of P (mg/L)] x [Volume of extractant (L) 4- mass of soil (kg)] 



Fractionation of Organic Phosphorus: 

Principles 

In general, the fractionation scheme follows the procedures developed by Bowman 
and Cole (1978) and modified by Sharpley and Smith (1985) and Ivanoff et al. (1998). 
Organic P in both calcareous and noncalcareous soils is fractionated into a labile pool, a 
moderately labile pool, and a nonlabile pool. The labile pool is extracted with 0.5M 
NaHC03 at pH 8.5. The extracted P includes both P„ and Pi in soil solution and sorbed 
on soil colloids. If desired, microbial biomass P in the soil can be determined at this 
point, via a chloroform (CHCI3) fumigation technique (Hedley and Stewart, 1982). The 
moderately labile pool is extracted with 1.0 M HC1, followed by 0.5 M NaOH. The 
NaOH extract is acidified with concentrated HC1 to separate the nonlabile fraction (humic 
acid fraction) from the moderately labile fraction (fulvic acid fraction). Finally, the 
highly resistant, nonlabile fraction is determined by ashing the residue from the NaOH 
extraction at 550°C for 1 h, followed by dissolution in 1.0 M sulfuric acid (H2SO4). The 
complete soil P fractionation scheme is shown in Figure 2. In all cases, P concentration 
in the extracts is determined colorimetrically by the phospho-molybdate method of 
Murphy and Riley (1962). Acid or alkaline extracts are neutralized prior to P 
determinations. Organic P in the extracts is calculated from the difference between total 
P and Pj. Total P in the extracts is measured after an aliquot is digested with 2.5 M 
H2SO4 and potassium persulfate (K2S2O8), according to the method of Bowman (1989), 
as modified by Thien and Myers (1992). 

Equipment 

1. Reciprocating shaker 

2. Centrifuge and 100 mL tubes 

3. Hot plate 

4. Muffle furnace 

5. Spectrophotometer 

Reagents 

1. 0.5 M sodium bicarbonate solution. Dissolve 42 g of NaHC03 in 1 L deionized 
water. Adjust the pH of this solution to 8.5 with 1 M NaOH (40 g of NaOH in 1 L 
deionized water). Avoid exposure of solution to air. Prepare fresh solution if 



54 



Methods for P Analysis, G.M. Pierzynski (ed) 

solution has been stored more than 1 month in a glass container. Solution can be 
stored >1 month in polyethylene, but pH should be checked each month. 

2. /?-nitrophenol indicator. Dissolve 0.25 g of p-nitrophenol in 100 mL of deionized 
water. 

3. 2 M and 1 M hydrochloric acid. Dilute 168 mL and 84 mL, respectively, of 
concentrated HC1 to 1 L with deionized water. 

4. Phospho-molybdate reagents. Dissolve 12 g of ammonium paramolybdate 
[(NH 4 )6M07 024'4H 2 0] in 250 mL of deionized water. Dissolve 0.2908 g of 
potassium antimony tartrate (KSbO C4H4O6) in 100 mL of deionized water. Add 
these solutions to 1 Lof 2.5 MH2SO4 (141 mL of concentrated H2SO4 diluted to 1 
L), mix thoroughly, and after cooling, dilute to 2 L with deionized water. Store 
solution (Reagent A) in a dark, cool place. To prepare reagent B, dissolve 1.056 g 
of L-ascorbic acid (CePLC^) in 200 mL of Reagent A, and mix. Reagent B should 
be prepared as needed, because it must be used within 24 h. 

5. 2.5 M sulfuric acid. Dilute 140 mL of concentrated H 2 SO4 to 1 L with deionized 
water. 

6. 2 M and 0.5 M sodium hydroxide. Dissolve 80 g and 20 g, respectively, of NaOH 
in 1 L deionized water. 

7 . Potassium persulfate (K 2 S 2 Og ) - reagent grade 

8. Chloroform (CHCI3 ) - ethanol free, reagent grade 

Procedures 
Labile Organic P: 

Weigh duplicate 1.0 g (oven-dry weight basis) samples of sieved (2 mm), field-moist 
soil into two 100 mL centrifuge tubes. To one tube, add 50 mL of 0.5 M NaHC03 and 
place sample horizontally on a reciprocating mechanical shaker for 16 h. At the end of 
the extraction period, centrifuge sample at 7000 rpm for 15 min and filter supernatant 
through Whatman No. 41 quantitative paper into a 50-mL volumetric flask. Bring to 
volume with deionized water and mix well. 

To determine labile Pi, transfer an aliquot containing 2 to 40 (ig P to a 50-mL 
volumetric flask, add five drops of p-nitrophenol indicator to the flask and adjust the pH 
with 2 M HC1 until the indicator color just changes from pale yellow to colorless. Add 
approximately 40 mL of deionized water to the flask, followed by 8 mL of Reagent B. 
Bring to volume with deionized water, and mix well. After 20 min., determine P 
concentration on a calibrated spectrophotometer at 880 nm. A blank containing the 0.5 M 
NaHC03 extracting solution should be analyzed with the sample. 

To determine total labile P in the extract, add 0.5 g of K 2 S 2 0g with a calibrated scoop 
to a 25-mL volumetric flask, transfer an appropriate aliquot (usually 1 to 5 mL, 
depending on P concentration) of the extract into the flask, and add 3 mL of 2.5 M 
H 2 SO4 . Digest sample on a hot plate at >150°C for 20 to 30 min. Digestion is complete 
after vigorous boiling subsides. Cool sample, add 5 mL of deionized water. After 
mixing, add five drops of p-nitrophenol indicator to the flask and adjust the pH with 5 M 
NaOH. Add approximately 10 mL of deionized water to the flask, followed by 4 mL of 
Reagent B. Bring to volume with deionized water, and mix well. After 20 min., 
determine P concentration on a calibrated spectrophotometer at 880 nm. 



55 



Methods for P Analysis, G.M. Pierzynski (ed) 



1.0 g (oven-dry basis) of 
moist soil in 100 mL tube 



50 mL 0.5M NaHC0 3 , 

shake 16 h, centrifuge, 

filter 



— Aliquots ► 
— Aliquots ► 



Labile P. 



Persulfate digestion <- 

I 



Total Labile P 



1.0 g (oven-dry basis) 

of moist soil in 100 

mL tube 



I 



2 mL CHC1 



(lyses microbial cells) 

1 



50 mL 0.5M NaHCO,, 

shake 16 h, centrifuge, 

filter 



1 



Labile P =Total Labile P- Labile P 

O 1 

Biomass P =Total Labile P- Total P 



50 mL 1.0M HC1 

shake 3 h, centrifuge, 

filter 



-Aiiquot> Moderately Labile P 

-Aliquot 



Persulfate digestion ► Total P 

Moderately Labile P a =Total P- P 



Rinse with deionized 
water 5 min, centrifuge 



► Discard Supernatant 



50 mL 0.5M NaOH, 

shake 3 h, centrifuge and 

filter 



— Aliquot - 



-Aliquot - 



Persulfate digestion 



Total P 



Acidfy to pH 0.2, -Aliquot- 
centrifuge 



Persulfate digestion 
4- 



Fulvic Acid P 



Humic Acid P = Total P- Fuvlic Acid P 



Rinse with deionized 
water 5 min, centrifuge 



Discard Supernatant 



Ash at 550°C, 1 h 



Dissolve in 1.0 M 
H 2 S0 4 , shake 24 h 



Nonlabile P 



Figure 2. Sequential fractionation scheme for organic P. 



56 



Methods for P Analysis, G.M. Pierzynski (ed) 

The difference between total labile P by persulfate oxidation and labile P ; gives an 
estimate of labile P„ . The Pi analysis should be performed as soon as possible after the 
soil extraction to minimize hydrolysis of P . 

To estimate P associated with soil microbial biomass, treat the second duplicate 
weighed sample with 2 mL of ethanol-free CHCI3 . Cover the uncapped tubes loosely 
with paper towels and place under a fume hood for 24 h. At the end of this period, 
extract samples with 0.5 M NaHC03 as previously described. The difference between the 
amounts of total labile P in the CHCI3 -treated and untreated duplicate soil samples 
determines biomass P that originated from lysed microbial cells. 

Moderately Labile Organic P: 

A two-step process is required to determine moderately labile P . Add 50 mL of 1 M 
HC1 to the residue from the labile P extraction and place sample on a reciprocating 
mechanical shaker for 3 h. An aliquot of 1 M HC1 should be used to rinse residue from 
filter paper used in the labile P extraction. After 3 h, centrifuge sample at 7000 rpm for 
15 min and filter supernatant through Whatman No. 41 quantitative paper into a 50-mL 
volumetric flask. Bring to volume with deionized water and mix well. Determine total P 
and Pi in the extract as previously described. Any P„ extracted in the 1 M HC1 is 
considered part of the moderately labile P fraction. 

Rinse the residue from the HC1 extraction with deionized water, shake for 5 min 
centrifuge, and discard the supernatant solution. Add 50 mL of 0.5 M NaOH to the 
residue and shake sample for 16 h. At the end of the extraction time, centrifuge sample at 
7000 rpm for 15 min. The supernatant contains both moderately labile P (fulvic acid P) 
and nonlabile P (humic acid P). To separate these fractions, remove an aliquot of the 
NaOH extract and acidify to pH 0.2 with concentrated HC1. At this pH, humic acids 
precipitate, and fulvic acids remain in solution. Centrifuge acidified sample at 7000 rpm 
for 15 min. Determine total P in both the NaOH extract and the acidified sample as 
previously described. Total P in the acidified sample is a measure of fulvic acid P. 
Estimate humic acid P by subtracting fulvic acid P from the total P measured in the 
NaOH extract (Figure 2). 

Nonlabile Organic P: 

To determine highly-resistant, nonlabile P , rinse the residue from the NaOH 
extraction with deionized water, shake for 5 min., centrifuge, and discard the supernatant 
solution. Place the residue in a crucible and ash at 550°C for 1 h. Dissolve ash by 
shaking in 1 M H2SO4 for 24 h, and measure P in solution as previously described. 

Calculations 

The amount of P in each fraction can be calculated using the following equation: 

P concentration is given fraction (mg/kg)= 

[Cone, of P (mg/L)] x [volume of extractant (L) 4- mass of soil (kg)] 
Comments: 

It should be remembered that P fractionation schemes are operationally defined. It is 
difficult to identify which discrete P compounds are extracted with each step. Moreover, 
hydrolysis of some P compounds by 1 M HC1 or 0.5 M NaOH, sorption of labile P, and 



57 



Methods for P Analysis, G.M. Pierzynski (ed) 

heterogeneity of soil particles within a sample may limit the accuracy of this fractionation 
procedure. 



References: 

Anderson, G. 1967. Nucleic acids, derivatives, and organic phosphates, p. 67-90. In: 

A.D. McLaren and G.H. Peterson (ed.). Soil Biochemistry. Marcel Dekker, New 

York. 
Bowman, R.A., and C.V. Cole. 1978. An exploratory method for fractionation of organic 

phosphorus from grassland soils. Soil Sci. 125:95-101. 
Bowman, RA. 1989. A sequential extraction procedure with concentrated sulfuric acid 

and dilute base for soil organic phosphorus. Soil. Sci. Soc. Am. J. 53:362-366. 
Chang, S.C., and M.L Jackson. 1957. Fractionation of soil phosphorus. Soil Sci. 84:133- 

144. 
Cross, A.F., and W.H. Schlesinger. 1995. A literature review of the Hedley 

fractionation: Applications to the biogeochemical cycle of soil phosphorus in natural 

ecosystems. Geoderma. 64:197-214. 
Fife, C.V. 1962. An evaluation of ammonium fluoride as a selective extractant for 

aluminum-bound soil phosphate: detailed study of soils. Soil Sci. 96:112-120. 
Halstead, R.L., and G. Anderson. 1970. Chromatographic fractionation of organic 

phosphates from alkali, acid, and aqueous acetylacetone extracts of soils. Can. J. Soil 

Sci. 50:11-119. 
Hedley, M.J., J.W.B. Stewart, and B.S. Chauhan. 1982. Changes in inorganic and 

organic soil phosphorus fractions induced by cultivation practices and by laboratory 

incubations. Soil Sci. Soc. Am. J. 46:970-976. 
Ivanoff, D.B., K.R. Reddy, and S. Robinson. 1998. Chemical fractionation of organic 

phosphorus in selected histosols. Soil Sci. 163: 36-45. 
Kuo, S. 1996. Phosphorus. In: D.L. Sparks (ed.) Methods of soil analysis. Agronomy 9. 

ASA-SSSA, Madison, WI. 
Murphy, J., and J. P. Riley. 1962. A modified single solution method for the 

determination of phosphate in natural waters. Anal Chem. Acta. 27:31-36. 
Peterson, G.W., and R.B. Corey. 1966. A modified Chang and Jackson procedure for 

routine fractionation of inorganic soil phosphates. Soil Sci. Soc. Amer. Proc. 30:563- 

565. 
Sharpley, A.N., and S.J. Smith. 1985. Fractionation of inorganic and organic phosphorus 

in virgin and cultivated soils. Soil Sci. Soc. Am. J. 49:127-130. 
Smillie, G.W., and J.K. Syers. 1972. Calcium chloride formation during extraction of 

calcareous soil with floride: II. Implications to the Bray-1 test. Soil Sci. Soc. Am. 

Proc. 36:25-30. 
Stott, D.E., and MA. Tabatabai. 1985. Identification of phospholipids in soils and 

sewage sludges by high performance liquid chromatography. J. Environ. Qual. 

14:107-110. 
Tate, K.R. 1984. The biological transformation of P in soil. Plant Soil. 76:245-256. 
Thien, S.J., and R. Myers. 1992. Determination of bioavailable phosphorus in soil. Soil 

Sci. Soc. Am. J. 56:814-818. 



58 



Methods for P Analysis, G.M. Pierzynski (ed) 



Williams, J.D.H., J.K.Syers, and T.W. Walker. 1967. Fractionation of soil inorganic 
phosphorus by a modification of Chang and Jackson's procedure. Soil Sci. Soc. 
Amer. Proc, 31:736-739. 



59 



Methods for P Analysis, G.M. Pierzynski (ed) 



Phosphorus Fractionation in Flooded Soils and Sediments 

Philip Moore, USDA-ARS, Fayetteville, AR 
Frank Coale, University of Maryland 

Introduction: 

Phosphorus (P) chemistry in soils and sediments is greatly influenced by the 
oxidation-reduction status (redox potential). Under oxidized conditions, ferric and 
manganic oxides and hydroxides are important adsorption sites for P. In addition, ferric 
and manganic phosphate minerals, such as strengite (FeP04 - 2H 2 0), and trivalent Mn 
phosphate (MnP03'1.5H 2 0) can form and persist under oxidized conditions. However, 
under reducing conditions these minerals are unstable, resulting in dissolution and release 
of soluble P into the soil solution (Patrick et al., 1973; Emerson, 1976; Emerson and 
Widmer, 1978; Boyle and Lindsay, 1986; Moore and Reddy, 1994). 

Since Fe and Mn phosphate mineral formation is controlled by the redox potential of 
the soil or sediment, it is important that soil and sediment samples that are collected 
under reduced conditions are handled appropriately during P fractionation to get an 
accurate picture of the P status. Allowing anaerobic sediments to become oxidized 
results in the rapid conversion of ferrous iron (Fe 2+ ) to ferric iron (Fe 3 + ). Within a very 
short time period (seconds to minutes), solid phase Fe(OH) 3 precipitates out of solution. 
Fresh ferric hydroxide precipitates have tremendous P sorption capacities, and they can 
cause the soluble P levels in the porewater to be reduced by orders of magnitude in 
minutes. To avoid this, samples should be maintained under anerobic conditions during 
the initial phases of P fractionation. 

Sequential extraction schemes for P (phosphorus fractionation) have been employed 
by various workers over the past 60 years, yet this is not an exacting science (Dean, 1938; 
Williams, 1950; Chang and Jackson, 1957; Williams et al, 1967; Chang et al, 1983). It 
must be kept in mind that these are rather crude methods, with many extractants causing 
the dissolution of more than one type of P solid phase. For example, sodium hydroxide is 
often used to extract Al and Fe-bound P (van Eck, 1982; Hieltjes and Lijklema, 1980). 
However, this compound will also extract organic P fractions, particularly in soils that 
have been heavily manured in the past. Hence, authors must be aware of the pitfalls and 
fallibility of the methods we are outlining, and use them only when they are the best 
procedure available. 

Materials: 

1 . PVC or plexiglas cylinder for taking cores 

2. Purified N 2 

3. Glove bag 

4. Vacumn pump 

5. Centrifuge and 250 mL centrifuge tubes with caps equipped with rubber septums 

Reagents: 

1. Deionized water 

2. 1MKC1 



60 



Methods for P Analysis, G.M. Pierzynski (ed) 

3. O.lMNaOH 

4. 0.5MHC1 

5. Concentrated HC1 (trace metal grade) 

Method: 

The P fractionation procedure described below is similar to that of van Eck (1982) as 
modified by Moore and Reddy (1994). 



Sampling: 

Flooded soil or sediment samples can be taken with a PVC or plexiglas cylinder. The 
coring device should be beveled from the outside so that it can be inserted into the 
sediment. Under certain conditions, such as in salt marshes or rice fields, it may be 
necessary to pound on the coring device with a hammer to reach the desired sampling 
depth. 

The sample can be returned to the lab in the sampling cylinder or it can transferred to a 
pre- weighed centrifuge tube. If it is to be transported in the cylinder, then a rubber 
stopper should be placed on the bottom of the core to hold the sediment in place. It is a 
good idea to tape the stopper in place. If the sample is taken under flooded conditions, 
leave some of the floodwater on top of the sample. If samples have been taken from a 
lake bottom, then the entire headspace should be filled with lake water and a stopper 
should be placed on the top of the cylinder as well. This reduces the amount of shaking 
and minimizes disturbance of the sediment/water interface. 

If samples are taken in flooded or saturated agricultural fields, transfer them directly 
into a 250 mL polycarbonate centrifuge tube. It is important that the sampling corer have 
an inside diameter slightly smaller than the inside diameter of the centrifuge tube. To 
take the sample, simply push the corer into the sediment to the desired depth (e.g., 10 
cm). It may be difficult to remove the core from the sediment without disturbing the 
sample. It may be necessary to hold the sediment in place from underneath the core 
(usually by hand) when pulling the core out of the ground to prevent the soil from falling 
out. If the soil is relatively fine textured the core can be rocked side to side and removed. 
Once the core has been removed from the sediment, pour the water off and place the 
cylinder over the mouth of the centrifuge tube. If the sample is from a coarse textured 
soil, it will fall into the tube. However, when clay contents are high, it will adhere to the 
sampling core. In this case it is necessary to have a ramrod with a rubber stopper (outside 
diameter slightly smaller than sampling cylinder' s inside diameter) to force the sample 
into the centrifuge tube. 

After the sample is in the centrifuge tube, tap the tube on a hard surface (palm of your 
hand) to allow any entrained air bubbles to escape to the surface. If these air bubbles are 
not removed, then the sample will become oxidized. 

Next, screw the lid onto the centrifuge tube and insert a 12 gauge needle through the 
rubber septum in the tube's top. Insert another 12 gauge needle that is connected via 
tygon tubing to the N 2 gas cylinder and begin purging. Purge the headspace for 5-10 
minutes with N2 at a pressure of about 10 psi. This pressure, coupled with the needle 
size, will result in a loud hissing sound; absence of the sound may mean the needle is 
clogged with sediment. Extra needles should be taken into the field in case this happens. 



61 



Methods for P Analysis, G.M. Pierzynski (ed) 

After purging the sample, remove the needle not connected to the N 2 first, then the other 
needle. This allows a positive pressure of N2 on the sample, so if the container leaks, the 
leak will be outward. 

If the samples are to be processed in less than two days, refrigeration is not required. 
For longer periods, the samples should be put on ice to slow down biological activity. It 
should be noted that many plastics, like polycarbonate, allow slow diffusion of oxygen. 
If samples are stored for months in the refrigerator, the sediment along the walls of the 
tubes will change color to red and orange, as oxygen enters the tube and oxidizes iron. If 
this happens, the sample should not be used. 

Water-Soluble P: 

The first fraction of P to be extracted from the sample is water-soluble P. If the 
sample was taken in intact sediment cores and the researcher desires to obtain a depth 
distribution of P in the core, then a glove bag is needed. Place the top of the core into the 
glove bag. Also place any supplies (spatula, purged centrifuge tubes, syringes, etc.) into 
the bag. Fill the bag with N2 gas, and empty it two or three times to make sure it is 
oxygen-free. Use a ramrod with rubber stopper (plunger) as described above to slowly 
push the sediment to the surface. Using the spatula, take the first sample to the desired 
depth [it helps to have the depth increment (e.g., 5 cm) marked on the plexiglas corer]. 
After the sediment has been placed in the tube, tap the tubes to get rid of bubbles. Then 
push the plunger upward another 5 cm (or whatever depth is desired). Repeat this 
process until all of the samples are in the tubes. 

Open the glove bag and purge the headspace in the centrifuge tube as described 
earlier. The headspace should be anerobic, if the glove bag worked correctly. However, 
trace quantities of O2 can cause problems, so this extra step is warranted. If the samples 
were transferred to centrifuge tubes in the field, purge them in the laboratory immediately 
prior to centrifugation to make certain the headspace is oxygen-free. 

First, record the weight of the tube plus sediment. Since the weight of the tube was 
recorded earlier, the wet sediment weight will be known. Centrifuge the samples at 7500 
rpm for 20 minutes. At this point the samples are most susceptible to oxidation, since the 
porewater is separated from the soil. Hence, do not open the centrifuge tubes unless you 
are ready to filter immediately. 

It is preferable to filter the samples quickly, so vacuum filtration is strongly 
recommended, using a 0.45 (im membrane filter. Turn on the pump and quickly open and 
pour the soil solution onto the filter. It should filter in a few seconds. Quickly pour the 
supernatant into a plastic sample container and acidify with concentrated HC1 to pH 2. It 
is mandatory that the water-soluble sample be acidified. Otherwise, when the sample 
oxidizes, soluble iron will precipitate soluble P, as discussed earlier. 

If pH measurements are to be taken, do not filter all of the sample. Using a 60 mL 
syringe, remove a suitable aliquot of the porewater for pH measurement. Hold the 
syringe upright and get rid of any air bubbles. Keep the sample in the syringe until pH is 
measured. Flooded soil/sediment samples have a high partial pressure of CO2 (often 
greater than 5%). If degassing occurs prior to pH measurement, the pH will often change 
by one to two units. 

The acidified, filtered sample for water-soluble P can be analyzed by several methods. 
If the Murphy-Riley method is used, then the anaylses can be referred to as soluble 



62 



Methods for P Analysis, G.M. Pierzynski (ed) 

reactive P. It is considered soluble since it passed through 0.45 (im membrane, and 
reactive since it reacted with the reagents in the Murphy-Riley method. 

The residual sediment from the water-soluble fraction will be used for the remaining 
fractionation. Hence, after the porewater has been removed, screw the lid back on the 
tube and purge with N2 to maintain anaerbic conditions. 

Loosely Sorbed P: 

Various salts have been used in the past for loosely sorbed P. van Eck (1982) utilized 
NH4CI for this purpose. However, in many studies focusing on P, it is also desirable to 
measure the amount of inorganic N present as ammonium. Hence, Moore and Reddy 
(1994) utilized KC1 for this fraction, so that exchangeable NH 4 (and exchangeable metals 
minus K) could be determined on one sample. 

After the porewater has been removed for water-soluble P, the tubes should be 
weighed to determine how much water was removed from the sample. Next, the tubes 
are placed into a glove bag and purged with N2 gas as described above. The sediment in 
the tubes should then be homogenized with a spatula, and a subsample (approximately 1 
gram dry weight) should be transferred into another pre-weighed centrifuge tube. 
Another subsample will be taken for moisture content, so that the exact weight of the 
sample for P fractionation is known. While still in the glove bag, add 20 mL of de- 
aerated 1 M KC1 to the tubes. When the tubes are removed from the glove bag, purge 
again with N2 gas to ensure the headspace is oxygen-free. 

Shake the tubes for 2 h on reciprocating shaker, then centrifuge at 7,500 rpm for 20 
minutes, and quickly filter through 0.45 (im filters as described above. The supernatant 
should be acidified to pH 2 with concentrated HC1. The sample can then be analyzed by 
the Murphy-Riley method. This fraction is loosely sorbed P. 

After this fraction has been taken, precautions to maintain anaerobic conditions are no 
longer needed. Decant excess KC1, then weigh again. Weights of each successive 
fraction are needed to calculate the entrained liquid (containing soluble P) from the prior 
extraction. 

Aluminum and Iron-bound P: 

The residual sediment from the KC1 extraction will be utilized "as is" for the next 
extraction (with NaOH). Add 20 mL of QAM NaOH to the sample, and shake for 17 
hours. Then centrifuge at 7500 rpm and filter through 0.45 (im membrane filters. 
Analyze using the Murphy-Riley method. This fraction is referred to as Al and Fe-bound 
P. 

It should be noted that some researchers will split this sample and digest half of the 
sample prior to analysis with Murphy-Riley. The difference between the undigested 
NaOH sample and the digested NaOH sample is referred to as "organic -bound P." 

Calcium-bound or Apatite P: 

After removing excess NaOH and weighing the previous sample, add 20 mL of 0.5 M 
HC1 and shake for 24 h. If the sediment contains free carbonates, open the samples 
during the first 15 minutes or so to relieve the pressure from CO2 buildup. After they 
have shaken for 24 hours, filter through 0.45 (im membrane filters, and analyze using the 



63 



Methods for P Analysis, G.M. Pierzynski (ed) 

Murphy-Riley method. This aliquot is referred to as Ca-bound P, but may also contain 
some organic P. 

Residual P: 

The remaining sample can then be analyzed for total P using a nitric-perchloric acid 
digestion or other suitable method. This is simply referred to as residual P, since it 
probably contains some Al and Fe-bound P, as well as organic P. Residual P can also be 
calculated by measuring total P on the original sample and subtracting the various 
fractions. 

References: 

Boyle, F.W., Jr. and W.L. Lindsay. 1986. Manganese phosphate equilibrium 

relationships in soils. Soil Sci. Soc. Am. J. 50:588-593. 
Chang, S.C. and M.L. Jackson. 1957. Fractionation of soil phosphorus. Soil Sci. 84:133- 

144. 
Chang, A.C., A.L. Page, F.H. Sutherland, and E. Grgurevic. 1983. Fractionation of 

phosphorus in sludge-affected soils. J. Environ. Qual. 12:286-290. 
Dean, L.A. 1938. An attempted fractionation of the soil phosphorus. J. Agric. Sci. 

28:234-246. 
Emerson, S. 1976. Early diagenesis in anaerobic lake sediments: Chemical equilibria in 

interstitial waters. Geochim. Cosmochim. Acta 40:925-934. 
Emerson, S. and G. Widmer. 1978. Early diagenesis in anaerobic lake sediments: II. 

Thermodynamic and kinetic factors controlling the formation of iron phosphates. 

Geochim. Cosmochim. Acta 42:1307-1316. 
Hieltjes, A.H.M and L. Lijklema. 1980. Fractionation of inorganic phosphates in 

calcareous sediments. J. Environ. Qual. 9:405-407. 
Moore, P. A., Jr. and K.R. Reddy. 1994. Role of Eh and pH on phosphorus geochemistry 

in sediments of Lake Okeechobee, Florida. J. Environ. Qual. 23:955-964. 
Murphy, J. and J.P. Riley. 1962. A modified single solution method for the 

determination of phosphate in natural waters. Analytica Chimica Acta 27:31-36. 
Patrick, W.H., Jr., S. Gotoh, and B.G. Williams. 1973. Strengite dissolution in flooded 

soils and sediments. Science 179:564-565. 
Williams, C.H. 1950. Studies on soil phosphorus: I. A method for the partial 

fractionation of soil phosphorus. J. Agric. Sci. 40:233-242. 
Williams, J.D.H., J.K. Syers, and T.W. Walker. 1967. Fractionation of soil inorganic 

phosphate by a modification of Chang and Jackson's procedure. Soil Sci. Soc. Am. 

Proc. 31:736-739. 
van Eck, G.T.M. 1982. Forms of phosphorus in particulate matter from the Hollands 

Diep/Haringvliet, the Netherlands. Hydrobiologia 91:665-681. 



64 



Methods for P Analysis, G.M. Pierzynski (ed) 

Determination of Phosphorus Retention and Flux in Soil 

Thanh H. Dao, USDA-ARS 

Introduction: 

In soils and sediments, physicochemical and biological processes act jointly to control 
the amount of phosphorus (P) that is in solution. The soluble reactive P fraction is taken 
up by plants, sequestered in soil, or disperses in the surrounding environment. Although 
the primary mechanism for environmental transport of P from agricultural soils is by 
erosion and surface runoff, specific instances of subsurface movement have been reported 
(Heckrath et al, 1995; Eghball et al, 1996; Gachter et al, 1998). Agricultural P inputs to 
nearby surface waters have been associated with toxic algal blooms and the depletion of 
oxygen in aquatic systems. An improved understanding of P retention and transport 
mechanisms is needed to develop management practices to mitigate P transport and 
inputs to surface waters. 

Typical methods used for assessing the environmental behavior of native and added P 
in terrestrial and aquatic ecosystems include procedures for measuring the retention 
capacity of soils and sediments and the associated kinetic parameters. Phosphorus 
retention has been commonly determined by batch equilibrium methods in which soil or 
sediment samples are agitated with P solutions of known concentrations (Graetz and 
Nair, 2000, this publication). The suspension is equilibrated for a sufficient time to 
achieve apparent equilibrium in the system. The advantages and disadvantages of the 
technique have been extensively reviewed (Green and Karickoff, 1990; Sparks et al., 
1996). 

Flow methods have also been used to study water and dissolved solute movement, the 
retention and desorption processes, for P in particular (Rao et al., 1979; van Riemsdijk 
and van der Linden, 1984; Miller et al., 1989; Beauchemin et al., 1996). Flow methods 
are open systems where solute and the reaction products with soil and sediment 
constituents are removed, minimizing re-adsorption, secondary precipitation reactions, or 
inhibition of desorption. Important parameters include water flux, chemical and 
hydrodynamic dispersion, sorption, exchange and desorption characteristics, and 
transformation rates coefficients. 

Applications: 

A flow displacement approach facilitates the simulation of the dynamic sorption- 
desorption, transformations, and transport of P in the soil and water system. 
Displacement studies provide insights in the kinetics of P release and physical and 
chemical non-equilibrium conditions that may influence nutrient mineralization and 
transport in soil. Columns experiments have been conducted to study the miscible 
displacement of organic chemicals (Green and Corey, 1971; Rao et al., 1979; Dao et al., 
1980; Wagenet and Rao, 1990) and for PO4-P in particular (van Riemsdijk and van der 
Linden, 1984; Miller et al, 1989; Chen et al, 1996). Breakthrough curves yield 
characteristics of the adsorption-desorption non-equilibrium and soil- solvent- solute 
interactions (Green and Karickoff, 1990; Chen et al., 1996). 



65 



Methods for P Analysis, G.M. Pierzynski (ed) 

Materials and Equipment: 

1. Columns made from stainless steel, glass, or PVC tubings of known inner diameter 
(ID) ranging from 5 to 100 mm and length ranging from 100 to 300 mm. 

2. A Mariott bottle setup to achieve a constant hydraulic head above column intake 
for steady-state flow. 

3. A fraction collector, operating on a time- or volume-based mode. 

4. A spectrophotometer for manual or automated P analysis. 

Reagents: 

1. A P-free nutrient solution. Deionized water or a 0.0 1M CaC12 solution. Dissolve 
1.47 g of CaC12. 2 H20 in deionized water and dilute to 1 L mark. 

2. A solution of known Br- concentration (10 mg Br/L). Dissolve .0149 g of KBr per 
L. 

3. A solution of known P concentration (10 mg P/L). Dissolve .056 g of K2HP04 
per L. 

4. A microbial growth inhibitor, such as acetone or chloroform (20 g/L of influent). 

Procedures: 

Soil/Sediment columns 

Either obtain intact soil cores or pack a column with uniformly mixed soil/sediment 
materials at overall density of 1.2-1.3 Mg m" 3 . The lower end of the column should be 
fitted with a fritted glass porous plate and a drainage port. To minimize mixing at the 
soil-porous plate interface, keep the pore size in the end-plate assembly as small as 
possible. 

P sorption 

Deliver from a Mariott bottle setup to achieve a constant hydraulic head above the 
column intake for steady-state flow. Collect effluent with a fraction collector. Acidify 
effluent fraction and analyze for P concentrations. 

P desorption 

Upon achieving a steady-state outflow P concentration, substitute a O.OlMCaCb 
solution, or P-free nutrient solution as the influent to study P desorption from the 
soil/sediment column. It is important to be able to switch rapidly from one solution to the 
other and minimize mixing of the two solutions at the influent assembly. Collect 
fractions of the effluent as previously, and analyze for P concentrations. 

Analysis ofP in effluent 

Filter effluent through a 0.45-(im membrane to remove any particulate matter, and 
acidify with HC1 (<pH 2). Determine phosphorus concentrations of effluent samples 
using spectrophotometric (Alpkem, 1994), inductively-coupled plasma atomic-emission 
spectroscopic (Soltanpour et al., 1979), or ion chromatographic (Nieto and 
Frankenberger, 1985) methods. 



66 



Methods for P Analysis, G.M. Pierzynski (ed) 

Calculations: 

Plot P concentrations against either time or volume of effluent to obtain an effluent or 
breakthrough curve (BTC). The analysis of BTCs is greatly facilitate by expressing P 
concentrations as relative or reduced concentration (C/C ) and the effluent volume as 
dimensionless pore volumes. Calculate the number of pore volume (V/V ) by dividing 
the amount of effluent by the liquid capacity of the column (V ). The latter can be 
calculated either as 

(i) V =ALq 

where: 

A = column cross-section area, 

L = length, and 

q = volumetric water content, 
or 

(ii) from the difference in the initial dry weight of the column and the weight of the 
saturated column at the end of the experiment. 

The retardation of P, R P h 0S , relative to the movement of the water front is the measure 
of interaction between soil and P. In simplest terms, the value of V/V„ at C/C = 0.5 is an 
approximation of R P hos- 

As pore geometry is unique for each soil column, a BTC for a non-reacting water 
tracer is also obtained, providing a reference R and a measure of pore water velocity. A 
potassium bromide (KBr) influent solution is used to obtain a Br" breakthrough curve. 
The ratio of R P h 0S to Rb r will yield the retardation factor for P. As needed, the sorption 
coefficient is determined from the following relationship between R and K when sorption 
is linearly related to solute solution-phase concentrations (e.g. at low solute 
concentrations), 



R = l + 



f r \ 



K«J 



K 



where: 

r = soil bulk density, and 
q = volumetric water content. 

Comments: 

The breakthrough of Br" is also determined in the effluent using potentiometric 
(Frankenberger et al., 1996) or ion-chromatographic method (Dao, 1991; Tabatabai and 
Frankenberger, 1996). Organic water tracers such as fluoro-benzoates have also been 
used in many water movement studies (Bowman, 1984). Multiple tracers can be used 
simultaneously, and relatively lower concentrations of tracers are needed as lower 
detection and quantification limits are attainable with liquid-chromatographic techniques. 

Graphical curve-fitting methods and numerical least-squares procedures are available 
to conveniently obtain estimates of retardation factor and dispersion coefficient for 
constant concentration and pulse-type effluent curves (van Genuchten, 1980; Parker and 
van Genuchten, 1984). Calculated effluent curves are based on an equation that 



67 



Methods for P Analysis, G.M. Pierzynski (ed) 



approximates closely the analytical solutions of the advective-dispersive transport 
equation (Danckwerts, 1953), 



1 



7 c =r rfc 



Rx-vt 

2(DRt) U2 



that, when x = L (column length) reduces to 



C„ 



-erfc 



AR 



V_ 

v n 



R- 



V_ 



where 



-VIYD, 



the Peclet number, P = 

R = retardation factor, 

v = pore water velocity, and 

erfc = the error function complement. 
The sum of squares of the residuals between observed and calculated effluent relative 
concentrations are minimized with iterative optimization of R and the Peclet number (or 
indirectly the dispersion coefficient D). 

Constant- volume solvent delivery pumps can be used for the metering of the influent 
solutions. Maintaining constant flow conditions is essential in displacement studies of 
extended duration. Programmable pumps can be used to study steady state or transient 
flow regimes. Transport studies under unsaturated conditions are performed by the 
inclusion of a vacuum chamber surrounding the column bottom and the fraction collector. 

References: 

Alpkem. 1994. Ortho-phosphate in soil extracts. In: Flow Solution HI Adv. Tech. Flow 

Anal., College Station, TX 
Beauchemin, S., R.R. Simard, and D. Cluis. 1996. P sorption-desorption kinetics of soil 

under contrasting land uses. J. Environ. Qual. 25:1317-1325. 
Bowman, R.S. 1984. Evaluation of some new tracers for soil water studies. Soil Sci. 

Soc. Am. J. 48:987-993. 
Chen, J.S., R.S. Mansell, P. Nkedi-Kizza, and BA. Burgoa. 1996. P transport during 

transient, unsaturated water flow in an acid sandy soil. Soil Sci. Soc. Am. J. 60:42- 

48. 
Danckwerts, P.V. 1953. Continuous flow systems. Chem. Engineer. Sci. 2:1-13. 
Dao, T.H., T. L. Lavy, and R. Sorensen. 1980. Atrazine degradation and residue 

distribution in soil. Soil Sci. Soc. Am. J. 43:1129-1134. 
Dao, T.H. 1991 Field decay of wheat straw and its effects on metribuzin and S-ethyl 

metribuzin sorption and elution from crop residues. J. Environ. Qual. 20:203-208. 
Eghball, B., CD. Binford, and D.D. Baltensperger. 1996. P movement and adsorption in 

a soil receiving long-term manure and fertilizer application. J. Environ. Qual. 

25:1339-1343. 



68 



Methods for P Analysis, G.M. Pierzynski (ed) 

Eghball, B., G.D. Binford, and D.D. Baltensperger. 1996. Phosphorus movement and 
adsorption in a soil receiving long-term manure and fertilizer application. J. Environ. 
Qual. 25:1339-1343 

Frankenberger, Jr., W.T., M.A. Tabatabai, D.C. Adriano, and H.E. Doner. 1996. 

Bromine, chlorine, and fluorine, p. 833-867. In: Sparks, D.L. (ed). Methods of soil 
analysis. Part 3. Chemical methods. Soil Sci. Soc. Am. Book Series No. 5. Soil Sci. 
Soc. Am., Madison, WI. 

Gachter, R., J.M. Ngatiah, and C. Stamm. 1998. Transport of phosphate from soil to 
surface waters by preferential flow. Environ. Sci. Tech. 32:1865-1869. 

Graetz, D.A., and V.D. Nair. 2000. Phosphorus sorption isotherm determination, p. 13- 
16 In: Pierzynski, G.M. (ed) Methods of P analysis for soils, sediments, residuals, 
and waters. SERA-IEG 17 South. Cooperative Series Bulletin No. 396, p. 35-38. 

Green, R.E., and J.C. Corey. 1971. Pesticide adsorption measurement by flow 

equilibration and subsequent displacement. Soil Sci. Soc. Am. Proc. 35:561-565. 

Green, R.E., and S.W. Karickhoff. 1990. Sorption estimates for modeling, p. 79-101. 
In: Cheng, H.H. (ed) Pesticides in the soil environment: Processes, impacts, and 
modeling. Soil Sci. Soc. Am. Book Series No. 2. Soil Sci. Soc. Am., Madison, WI. 

Heckrath, G., P.C. Brooks, P.R. Poulton, K.W.T. Goulding. 1995. Phosphorus leaching 
from soils containing different P concentrations in the Broadbalk Experiment. J. 
Environ. Qual. 24:904-910. 

Miller, D.M., M.E. Sumner, and W.P. Miller. 1989. A comparison of batch and flow- 
generated anion adsorption isotherms. Soil Sci. Soc. Am. J. 53:373-380. 

Nieto, K.F., and W.T. Frankenberger, Jr. 1985. Single ion chromatography: I. Analysis 
of inorganic anions in soils. Soil Sci. Soc. Am. J. 49:587-592. 

Parker, J.C, and M. Th. Van Genuchten. 1984. Determining transport parameters from 
laboratory and field tracer experiments. VA Agric. Exp. Stn. Bull. 84. 

Rao, P.S.C., J.M. Davidson, R.E. Jessup, and H.M. Selim. 1979. Evaluation of 
conceptual models for describing kinetics of adsorption-desorption of pesticides 
during steady-state flow in soils. Soil Sci. Soc. Am. J. 43:22-28. 

Soltanpour, P.N., S.M. Workman, and A.P. Schwab. 1979. Use of inductively coupled 
plasma spectrometry for the simultaneous determination of macro- and micronutrients 
in NH4HCO3-DPTA extracts of soils. Soil Sci. Soc. Am. J. 43:75-78. 

Sparks, D.L., S.C. Fendorf, C.V. Toner IV, and T.H. Carski. 1996. Kinetic methods and 
measurements, p. 1275-1307. In: Sparks, D.L. (ed). Methods of soil analysis. Part 
3. Chemical methods. Soil Sci. Soc. Am. Book Series No. 5. Soil Sci. Soc. Am., 
Madison, WI. 

Tabatabai, M. A., and W.T. Frankenberger, Jr. 1996. Liquid chromatography, p. 225- 
245. In: Sparks, D.L. (ed). Methods of soil analysis. Part 3. Chemical methods. 
Soil Sci. Soc. Am. Book Series No. 5. Soil Sci. Soc. Am., Madison, WI. 

van Genuchten, M. Th. 1980. Determining transport parameters from solute 
displacement experiments. Res. Rep. 118 US Salinity Lab., Riverside, CA. 

van Riemsdijk, V.H., and A.M.A. van der Linden, 1984. Phosphate sorption in soils: U. 
Sorption measurement technique. Soil Sci. Soc. Am. J. 48:541-544. 

Wagenet, R. J., and P.S.C. Rao. 1990. Modeling pesticide fate in soils, p. 351-399. In: 
Cheng, H.H. (ed) Pesticides in the soil environment: Processes, impacts, and 
modeling. Soil Sci. Soc. Am. Book Series No. 2. Soil Sci. Soc. Am., Madison, WI. 



69 



Methods for P Analysis, G.M. Pierzynski (ed) 



Residual Materials 



70 



Methods for P Analysis, G.M. Pierzynski (ed) 

Sampling Techniques for Nutrient Analysis of Animal 
Manures 

D.A. Crouse, S.C. Hodges, C.R. Campbell, J.P. Zublena, North Carolina State 
University 



Introduction: 

Nutrient concentrations vary in most wastes. A review of samples analyzed by the 
North Carolina Department of Agriculture and Consumer Services Agronomic Division 
showed the available nitrogen in animal waste varies greatly. For example, in swine 
lagoon liquids, nitrogen can range from 3 to 73 mg/L, in dairy slurry the range is 12 to 
30,000 mg/L, and in a lagoon on a poultry operation with a liquid waste management 
system the range is 12 to 39,000 mg/L. This is a broad range of nutrient levels with the 
maximum and minimum values differing by more than a hundredfold. These numbers 
should send a clear message to users of animal waste: Average nutrient estimates may be 
suitable for the purposes of developing a waste management plan, but these averages are 
not adequate for calculating proper application rates. 

Proper sampling is the key to reliable waste analysis. No analytical method, statistical 
calculation or laboratory quality control program can generate meaningful data from a 
poorly representative sample. If the waste product to be analyzed is entirely homogenous, 
then a single sample, no matter how small in weight or volume or where it is taken, 
would be completely representative of the product (Chai, 1996). But, since animal wastes 
are inherently heterogeneous, proper sampling techniques are critically important. 
Reliable samples typically consist of material collected from a number of locations 
around the lagoon or waste storage structure. The sampling methodology described 
herein has been adapted from a North Carolina Cooperative Extension Publication - 
Waste Analysis (Zublena and Campbell, 1993) developed to educate farmers on the 
proper techniques for waste sampling. The North Carolina Department of Environment 
and Natural Resources has adopted the procedures as guidance for sampling to meet 
monitoring conditions in the permits issued to confined animal feeding operations. 

Obviously, sampling methods vary 
according to the type of waste. This 
publication will address liquid wastes 
and solids. The liquid waste section 
will address lagoon liquid (effluent) 
and slurries. The solid waste section 
will address waste products such as 
dairy dry stacks and poultry litter. 



Liquid Wastes: 

Lagoon Liquid 

Premixing the surface liquid in the 
lagoon is not needed, provided it is 
the only waste component that is 
being pumped for land application. 



Wooden pole (10 feet) 




Plastic cup 



Figure 1. Liquid waste sampling device. 



71 



Methods for P Analysis, G.M. Pierzynski (ed) 

Farms where multistage lagoon systems exist should have the samples collected from the 
lagoon they intend to pump for crop irrigation. 

Samples should be collected using a clean, plastic container similar to the one shown 
in Figure 1 . Galvanized containers should never be used for collection, mixing, or 
storage due to the risk of contamination from metals, such as Zn. A 500 mL sample of 
material should be taken from at least eight sites around the lagoon and then mixed in the 
larger clean, plastic container. Waste should be collected at least 2 m from the edge of the 
lagoon at a depth equivalent to that of the irrigation intake line in the lagoon, usually 
about 15 cm deep. Floating debris and scum should be avoided. A 500-mL subsample of 
the mixed material should be sent to the laboratory. 



Clean-out dowel 

(1 inch diameter 

PVC pipe) 



PVCpipo 

(2 inches diameter, 

6 feet long) 




Rubber bail 
{2 1 /; ince* diameter) 



Figure 2. Composite sampling device. 



Liquid Slurry 

Waste materials applied as a 
slurry from a pit or storage pond 
should be mixed prior to 
sampling. If mixing occurs prior 
to sampling, the liquid sampling 
device pictured in Figure 1 can be 
used. If a storage structure 
without agitation is sampled, use 
the composite sampling device as 
shown in Figure 2. Waste should 
be collected from approximately 
eight areas around the pit or pond 
and mixed thoroughly in a clean, 
plastic container. A 6-foot section 
of 1- to 2-inch plastic pipe can 
also be used: Extend the pipe into the pit; pull up the ball plug (or press your thumb over 
the end to form an air lock); remove the pipe from the waste; and release the air lock to 
deposit the waste in the plastic container. 

Collect about a 500-mL subsample in a clean plastic container for transport to the 
laboratory for analysis. The sample should not be rinsed into the container, since doing so 
skews the measured nutrient analysis relative to the analysis of the actual collected 
sample. However, if water is typically added to the waste prior to land application to aid 
in agitation and pumping, a proportionate quantity of water should be added to the 
collected sample prior to analysis. 

Whether sampling lagoon liquids or slurries, certain procedures are similar. All liquid 
waste samples collected and submitted for analysis should be placed in a sealed, clean, 
plastic container for storage and transport to the laboratory. Glass is not recommended 
due to potential damage to the container during transport. Samples should be tightly 
sealed as soon as possible. Some headspace should be left in the container to allow for 
some expansion of gases, lowering the potential for the container to rapidly erupt when 
opened in the laboratory. However, headspace should not exceed 2.5cm in order to 
minimize the potential for off-gassing of ammonia from solution. Samples that cannot be 
shipped on the day they are collected should be refrigerated. The most frequent changes 
in waste samples, be it solid liquid or sludge, are volatile losses, biodegradation, 
oxidation and reduction. Low temperatures reduce biodegradation and sometimes volatile 



72 



Methods for P Analysis, G.M. Pierzynski (ed) 

losses, but freezing liquid samples can cause degassing (Bone, 1988). Anaerobic samples 
must not be exposed to air for significant periods of time. 

Solid Wastes 

Dry Stacks 

Solid waste samples should represent the average moisture content of the waste. A 500 
cm sample is recommended. Samples should be taken from approximately eight 
different areas in the waste, placed in a clean, plastic container, and thoroughly mixed. 
Approximately 500 cm of the mixed sample should be placed in a plastic bag, sealed, 
and analyzed as soon as possible. Samples stored for more than two days should be 
refrigerated. Figure 3 shows a device for sampling solid waste. 



Poultry Litter 

If collecting poultry litter from 
a stockpile or dry litter storage 
shed, follow the procedure for 
Dry Stacks. If sampling directly 
from the house, samples should be 
taken from approximately 20 to 
30 different areas in the house. 
The samples should be placed in a 
clean, plastic container and 
thoroughly mixed. When 
sampling, be careful to get a 
representative sample. The 
number of samples taken from 
around the waterers, feeders, and 



Dowel 



Metal rod 




Plastic Container 
(5 gallons) 



Ttifn-walted metal 

tubing 
(1 inch diameter) 



Figure 3. Solid-waste sampling device. 



brooders should be proportionate to the area occupied by each. Sample only to the depth 
the house will be cleaned, avoiding collecting soil from underneath the litter. Litter from 
broiler breeder houses should be sampled after the slats are removed and the manure and 
litter have been mixed. Approximately 500 cm 3 of the mixed sample should be placed in 
a plastic bag, sealed, and analyzed as soon as possible. Samples stored for more than two 
days should be refrigerated. 

References: 

Bone, L.T. 1988. Preservation techniques for samples of solids, sludges and non-aqueous 

liquids, p. 409 In L.H. Keith (ed.) Principals of environmental sampling. Am.. Chem.. 

Soc. Washington, D.C. 
Chai, E.Y., 1996. A systematic approach to representative sampling in the environment. 

p. 33-44. In J.H. Morgan (ed) Sampling environmental media, ASTM STP 1282. Am. 

Soc. For Testing and Materials. 
Zublena, J.P. and C.R. Campbell. 1993. SoilFacts: Waste analysis. North Carolina 

Cooperative Extension publication series. AG-439-33. 



73 



Methods for P Analysis, G.M. Pierzynski (ed) 



Determining Water-Soluble Phosphorus in Animal Manure 

M.L. Self-Davis, University of Arkansas 
P.A. Moore, Jr., USDA-ARS, Fayetteville, AR 



Introduction: 

There are no "standard methods" for many of the tests dealing with solid animal 
wastes. Therefore, the analyses of animal waste are usually modifications of "standard 
methods" for other substances (Overcash et al., 1975). The procedure for determining 
water-soluble P in animal manure is a modification of a method used to determine water- 
soluble P in soils (Olsen and Sommers, 1982). This method was originally developed for 
soils to examine the chemical composition of the soil solution that surrounds plant roots 
(Adams, 1974). The modification of this method presented here was utilized by Moore 
and Miller (1994). 

Sampling: 

The composition of animal manure varies greatly with location in the production 
facilities. To adequately describe the chemical and/or microbial composition of the 
manure, proper sampling techniques are needed. The following example explains how to 
obtain representative samples: (1) Divide the production facility to be sampled into three 
zones. If the buildings run in an east- west direction, then divide them into the northern 
third, middle third, and southern third. (2) Start in one zone and, while walking down the 
length of the building in a zigzag pattern, take about 10-15 samples, and place them in a 
plastic bucket. Note: For dryer materials (poultry litter), a soil probe works well. Due to 
the consistency of manure in dairy or swine facilities, a small shovel is more 
appropriate. (3) Make sure that if sampling inside a production facility, some of the 
sample (a representative portion) comes from under the feeders and waterers. The 
sample should be taken from the surface to just above the floor (until the resistance of the 
manure does not allow you to easily push the sampler in). (4) Mix the contents of the 
bucket well and pour about 100 g of the sample into a labeled freezer bag or plastic 
container. (5) Repeat this process in the other two zones of the facility. 

Equipment: 

1. Shaker (reciprocating or end-over-end) 

2. Centrifuge 

3. Centrifuge tubes (250 mL) 

4. Filtration apparatus (0.45-(im pore diameter) 

5. Spectrophotometer with infrared phototube for use at 880 nm 

6. Acid-washed glassware and plastic bottles: graduated cylinders (5 mL to 100 mL), 
volumetric flasks (100 mL, 500 mL, and 1000 mL), storage bottles, pipets, dropper 
bottles, and test tubes or flasks for reading sample absorbance. 

Reagents: 

1. Concentrated hydrochloric acid (HC1) 



74 



Methods for P Analysis, G.M. Pierzynski (ed) 

2. Reagents used for ascorbic acid technique, Murphy-Riley (1962) 

Procedure: 

Weigh 20 g of fresh manure into a 250 mL centrifuge tube. Manure is not as 
homogeneous as soils. Therefore, a large sample is needed to get a good representation 
of the material. Add 200 mL of distilled water and shake for two hours. This ratio of 20 
g manure to 200 mL distilled water leaves sufficient room in the centrifuge tube for 
proper shaking. Centrifuge at 6,000 rpm for 20 minutes. Filter the solution through a 
0.45 (im membrane filter. Acidify to pH 2.0 with HC1 to prevent precipitation of 
phosphate compounds (normally add about 5 drops of concentrated HC1 per 20 mL). 
Freeze the sample if it is not going to be analyzed that day. Previous articles discussing 
the colorimetric determination of P have noted that hydrolysis of condensed phosphates 
can occur when the solution is acidified or in contact with acid for extended periods of 
time (Lee et al., 1965). Also, at this pH level, there is the possibility of flocculation of 
organics. However, it is necessary to make the sample solution as stable as possible, 
especially when there is a delay between the extraction process and actual analysis. It is 
vital to ensure that P remains in solution. Therefore, the negative effects of acid addition 
are often considered minimal. 

In order to calculate the amount of soluble P per kilogram of dry manure, the water 
content of the manure should be measured. On the same day the manure is extracted, 
weigh out another subsample (approximately 10 g) into a pre-weighed metal container 
and dry in a forced draft oven at 60°C for 48 hours. 

Analysis: 

For determining water-soluble P in animal manure, analyze the samples with a 
Technicon Auto- Analyzer (Technicon 1976) using the Murphy-Riley method (1962). 
Since this method does not quantify all the P in solution, it is referred to as "reactive" P. 
Anything that passes through a 0.45 (im filter is referred to as "soluble" P. Hence with 
this method you are determining soluble reactive P (SRP). This form of P is the most 
available for uptake by algae and higher plants. 

Other methods of P analysis can be used. Any spectrophotometer with an infrared 
phototube for use at 880 nm can be used. Also, samples can be analyzed using an 
inductively coupled plasma-atomic emission spectrometry (ICP-AES) which will also 
measure dissolved P. Therefore filtered samples are used to determine total dissolved P 
(TDP) with ICP-AES. 

Calculations: 

It is preferred to report P concentrations on a dry weight basis (mg P/kg dry manure) 

Manure P cone, (mg/kg) = 

[P cone, in extract (mg/L)] x [Extractant volume (L) 4- Mass of dry manure (kg)] 

If presenting on an "as is" basis: 

Manure P cone, (wet basis) (mg/kg) = 

[P cone, in extract (mg/L)] x [Extractant volume (L) 4- Mass of wet manure (kg)] 



75 



Methods for P Analysis, G.M. Pierzynski (ed) 

Comments: 

It can be difficult to filter manure extracts (particularly swine and dairy manure). To 
improve the filter process first try increasing the centrifuge speed from 6,000 to 8,000 
rpm or higher (be sure to note the maximum rpm your centrifuge tubes can withstand). 
Also, samples can be prefiltered though a glass fiber filter to prepare them for 0.45 (im 
membrane filtration. If filtering is still difficult, manure-to-water ratios can be increased 
(from 1:10 to 1:15 or 1:20). 

References: 

Adams, F. 1974. Soil solution. P. 441-481. In E.W. Carson (ed.) The plant root and its 

environment. University Press of Virgnia, Charlottesville. 
Lee, G.F., N.L. Clesceri, and G.P. Fitzgerald. 1965. Studies on the analysis of phosphates 

in algal cultures. Internat. J. Air Water Poll. 9:715-722. 
Moore, PA., Jr., and D.M. Miller. 1994. Decreasing phosphorus solubility in poultry 

litter aluminum, calcium, and iron amendments. J. Environ. Qual. 23:325-330. 
Murphy, J., and J. P. Riley. 1962. A modified single solution method for the 

determination of phosphate in natural waters. Anal. Chem. Acta 27:31-36. 
Olsen, S.R. and L.E. Sommers. 1982. Phosphorus, p. 403-430 In A.L. Page et al. (Ed.) 

Methods of soil analysis. Part 2. 2nd ed. Agronomy Monogr. 9. ASA and SSSA, 

Madison, WI. 
Overcash, M.R., A.B., Hashimoto, D.L. Reddell, and D.L. Day. 1975. Evaluation of 

chemical analysis for animal wastes. In Standardizing Properties and Analytical 

Methods Related to Animal Waste Research. ASAE SP-0275, ASAE, St Joseph, MI, 

pp. 335-55. 
Technicon. 1976. Individual/simultaneous determination of nitrogen and/or phosphorus 

in BD digest. Industrial method no. 329-74W/A. Technicon Industrial Systems, 

Tarrytown, NY. 



76 



Methods for P Analysis, G.M. Pierzynski (ed) 

Total Phosphorous in Residual Materials 
M.R. Bender and C.W. Wood, Auburn University 



Introduction: 

A review of literature pertaining to the analysis of total P in residual materials shows 
the use of varied methods. Many methods employed are the same as those used in the 
determination of total P in soil, such as NaHCCh fusion and alkali oxidation as described 
by Olsen and Sommers (1982) and Dick and Tabatabai (1977), respectively (Wen et al., 
1997; Harris et al., 1994). In this case, it is important that the selected method effectively 
oxidizes the organic matter of the residual material, since this component may contain P. 

The methods discussed here are perchloric acid digestion, nitric acid-sulfuric acid 
digestion, and persulfate oxidation used in conjunction with colormetric methods for 
determination of total P as described by APHA (1989), and a rapid perchloric acid 
digestion for analysis of total P by ion chromatography developed by Adler (1995). These 
methods have been developed for the organic materials found in wastewater and other 
types of residual materials. 

Perchloric Acid Digestion (APHA (1989)): 

Reagents 

1. Concentrated HN0 3 

2. 70-72% HCIO4 reagent grade 

3. 6MNaOH 

4. Methyl orange indicator solution 

5. Phenolphthalein indicator aqueous solution 

Procedure 

Add a known volume of a well-mixed sample to a 125 mL Erlenmeyer flask, and 
acidify to a methyl orange endpoint (from orange to red) with concentrated HNO3 . Add 5 
mL more of HNO3. Evaporate solution to 15 to 20 mL on a steam bath or hot plate. Add 
10 mL each of concentrated HNO3 and HCIO4 to the flask. Be sure to cool the flask 
before each addition. After adding a few boiling chips, heat flask on a hot plate, and 
evaporate until dense, white fumes of HCIO4 appear. If the solution is not clear, cover 
the flask with a watch glass and keep solution barely boiling until it clears. If necessary, 
10 mL more of concentrated HNO3 can be added to aid oxidation. Cool the digested 
solution, and add 1 drop of phenolphthalein indicator solution. Then add 6 M NaOH until 
solution turns pink in color. If necessary, filter the neutralized solution to remove 
particulate material. Wash the filter liberally with distilled water. Bring the volume of the 
solution to 100 mL with distilled water. 

To determine total P, use one of the colormetric methods discussed in the colormetric 
methods section of this chapter. Please note that choice of colormetric method depends 
on the concentration range of orthophosphate in the sample. The 
vanadomolybdophosphoric acid method can be used for samples that range between 1 to 
20 mg P/L. The ascorbic acid method can be used for samples that range between 0.01 to 
6 mg P/L. 



77 



Methods for P Analysis, G.M. Pierzynski (ed) 

Comments 

The perchloric digestion method is recommended for samples that are difficult to 
digest. 

Caution must be taken when mixing HCIO4 with organic materials. To avoid a violent 
reaction: 

1 . Do not add HCIO4 to a hot solution containing organic matter. 

2. Begin digestion of sample containing organic material with HNO3 first, then 
complete digestion with HNO3 and HCIO4 mixture. 

3. Only use a fume hood designed for HCIO4 use. 

4. Do not allow solution to evaporate to dryness. 

Nitric Acid and Sulfuric Acid Digestion (APHA (1989)): 

Reagents 

1. Concentrated H2SO4 

2. Concentrated HNO3 

3. Phenolphthalein indicator aqueous solution 

4. lMNaOH 

Procedure 

Add a known volume of a well-mixed sample to a micro-kjeldahl flask. Add 1 mL of 
concentrated H2SO4 and 5 mL of concentrated HNO3. Digest the solution to a volume of 
1 mL, and then continue digestion until solution becomes colorless to remove HNO3. 
Cool solution, then add 20 mL of distilled water. Add 1 drop of phenolphthalein indicator 
solution and add 1 M NaOH to the solution until a faint pink color is reached. If 
necessary, filter neutralized solution to remove particulate material. Wash filter liberally 
with distilled water. Bring the volume of the solution to 100 mL with distilled water. 

To determine total P use one of the colormetric methods discussed in the colormetric 
methods section of this chapter. Please note that choice of colormetric method depends 
on the concentration range of orthophosphate in sample. The vanadomolybdophosphoric 
acid method can be used for samples that range between 1 to 20 mg P/L. The ascorbic 
acid method can be used for samples that range between 0.01 to 6 mg P/L. 

Comments 

Nitric acid and sulfuric acid digestion is recommended for most samples. 



Persulfate Oxidation Method (APHA (1989)): 

Reagents 

1 . Sulfuric acid solution (H 2 SO4 ) . Prepare by adding 300 mL of concentrated H 2 SO4 
to 600 mL of distilled water. Dilute solution to 1 L with distilled water. 

2. Ammonium persulfate ((NH 4 ) 2 S 2 0% ) solid or potassium persulfate (K 2 S 2 Og ) solid 

3. lM(NaOH) 

4. Phenolphthalein indicator aqueous solution 



78 



Methods for P Analysis, G.M. Pierzynski (ed) 



Procedure 

Add 50 mL of a well-mixed sample (or any other suitable volume) to a flask. Add 1 
drop of phenolphthalein indicator solution. If a red color develops, add H2SO4 solution 
dropwise until color disappears. Then add 1 mL of H2SO4 solution and either 0.4 g of 
(NH 4 ) 2 S2 08 or 0.5 g of K1S2O8. Boil the solution gently on a preheated hot plate for 30 
to 40 min or until a final volume of 10 mL is reached. Those samples containing 
organophosphorous may take as much as 1.5 to 2 hr for complete digestion. Cool solution 
and dilute to 30 mL with distilled water. Add 1 drop of phenolphthalein indicator 
solution. Then add 1 M NaOH until solution turns a faint pink color. Heat the solution for 
30 min in an autoclave or a pressure cooker at 98 to 137 kPa, then cool the solution. Add 
1 drop of phenolphthalein indicator solution. Then add 1 M NaOH until solution turns a 
faint pink color. Bring the volume of the sample to 100 mL with distilled water. If a 
precipitate forms, do not filter. The precipitate will redissolve during the colormetric 
method used to determine total P. Mix solution well before further subdivision of the 
sample. 

To determine total P use one of the colormetric methods discussed in the colormetric 
methods section of this chapter. Please note that choice of colormetric method depends 
on the concentration range of orthophosphate in sample. The vanadomolybdophosphoric 
acid method can be used for samples that range between 1 to 20 mg P/L. The ascorbic 
acid method can be used for samples that range between 0.01 to 6 mg P/L. 

Comments 

Though the persulfate digestion method is a simple method, it may be prudent to 
check this method against one of the other methods described in this chapter. 

COLORIMETRIC METHODS 

Vanadomolybdophosphoric Acid Method: 

Reagents 

1 . Phenolphthalein indicator aqueous solution 

2. 6 M HC 1 or similar strength solution of H 2 S0 4 or HNO3 . 

3. Activated carbon (Darco G60 or equivalent). Rinse with distilled water to remove 
fine particulate material. 

4. Vanadate-molybdate reagent. Prepare solution A by dissolving 25 g ammonium 
molybdate ((NHO6M07O24 4H 2 0) in 300 mL of distilled water. Prepare solution B 
by dissolving 2.5 g of ammonium metavanadate (NH4VO3) by heating to boil in 
300 mL of distilled water. Cool solution and then add 330 mL cone. HC1. Cool 
solution B to room temperature. Pour solution A into solution B, mix, and then 
dilute to 1 L with distilled water. 

5. Standard P solution. Prepare by dissolving 219.5 mg of anhydrous KH2PO4 in 
distilled water. Dilute solution to 1 L with distilled water. (1.00 mL = 50.00 (ig 
PO4-P). 



79 



Methods for P Analysis, G.M. Pierzynski (ed) 

Procedure 

If sample pH is greater than 10, add 1 drop of phenolphthalein indicator solution to 50 
mL sample and add 6 MHO drop until the indicator changes color. Dilute sample to 100 
mL. To remove excess color, shake sample with 200 mg of activated carbon for 5 min in 
an Erlenmeyer flask. Place 35 mL or less of sample in a 50 mL volumetric flask. Add 10 
mL of vanadate-molybdate solution to the flask and dilute the contents to 50 mL with 
distilled water. To prepare a blank, add 35 mL of distilled water to a 50 mL volumetric 
flask in place of sample. Prepare a standard curve by using suitable volumes of standard 
solution in place of sample. Add standard solution to a 50 mL volumetric flask. Add 10 
mL of vanadate-molybdate solution to the flask and dilute the contents to 50 mL with 
distilled water. After 10 min or more read absorbance of sample against blank. For 
solutions with 1-5 mg P/L, 2-10 mg P/L, or 4-18 mg P/L measure absorbence at 400, 420, 
or 470 nm, respectively. 

Calculations 

To calculate mg P/L: 

mg P/L = [mg P (in 50 mL final volume) x 1000] ^[sample volume (mL)] 

Comments 

Check activated carbon for P. Phosphorus in the activated carbon can result in high 
reagent blanks. 

Use acid- washed glassware for determining low concentrations of P. Wash glassware 
with a P-free detergent, then clean all glassware with hot, diluted HC1 and rinse well with 
distilled water. For a P range of 1.0 to 5.0 mg P/L use a filter wavelength of 400 nm for 
the spectrophotometer. For a range of 2.0 to 10 mg P/L use a filter wavelength of 420 nm 
for the spectrophotometer. For a P range of 4.0 to 18 mg P/L use a filter wavelength of 
470 nm for the spectrophotometer. 

Ascorbic Acid Method: 

Reagents 

1. 2.5 M sulfuric acid (H2SO4). Prepare by diluting 5 mL of concentrated H2SO4 into 
500 mL of distilled water. 

2. Potassium antimonyl tartrate solution (K(SbO)C4H 4 06 1/2FFO). Prepare by 
dissolving 1.3715 g K(SbO)C 4 H4 6 1/2H 2 in 400 mL of distilled water in a 500 
mL volumetric flask, and dilute to volume with distilled water. Store reagent in a 
glass- stoppered bottle. 

3. Ammonium molybdate solution ((NH 4 ) 6 Mo7 02 4 4H 2 0). Prepare by dissolving 20 
g (NH 4 ) 6 Mo7 02 4 4H 2 in 500 mL of distilled water. Store reagent in a glass- 
stoppered bottle. 

4. 0.0 1M ascorbic acid. Prepare by dissolving 1.76 g of ascorbic acid in 100 mL of 
distilled water. Reagent is stable for approximately 1 week at 4°C. 

5. Mixed reagent. Prepare by mixing 50 mL 5N FFSO4, 5 mL potassium antimonyl 
tartrate solution, 15 mL ammonium molybdate solution, and 30 mL ascorbic acid 
solution. Mix after addition of each reagent. Be sure that all reagents are at room 
temperature before mixing, and be sure to mix in the order given. If turbidity 



80 



Methods for P Analysis, G.M. Pierzynski (ed) 

forms during the combination of reagents, shake and allow to stand until turbidity 
disappears before continuing. 

6. Stock P solution. Prepare by dissolving 219.5 mg of anhydrous KH2PO4 in 
distilled water. Dilute solution to 1 L with distilled water. (1.00 mL = 50.00 (ig 
PO4-P). 

7. Standard P solution. Prepare by diluting 50 mL of stock P solution to 1000 mL of 
distilled. (1.00 mL = 2.50 (Xg P0 4 -P). 

Procedures 

Pipet 50 mL of sample into a clean, dry test tube or a 125 mL Erlenmeyer flask. Add 
1 drop of phenolphthalein indicator solution, if a pink color develops add 2.5M H2SO4 
dropwise to the solution. Add 8.0 mL of mixed reagent to the solution and mix 
thoroughly. Prepare a standard curve by using suitable volumes of standard solution in 
place of sample. Use a series of 6 standard solutions within the approximate range of 0.01 
to 2.0 mg P/L. After 10 min and before 30 min measure absorbance at 880 nm. Use a 
reagent blank as a reference solution. 

Calculations 

To calculate mg P/L: 

mg P/L = [ mg P (in approximately 58 mL final volume) x 1000] -=- [sample volume 
(mL)] 

Comments 

For a P range of 0.30 to 2.0 mg P/L use a light path of 0.5 cm for the spectro- 
photometer. For a range of 0.15 to 1.3 mg P/L use a light path of 1.0 cm for the 
spectrophotometer. For a range of 0.01 to 0.25 mg P/L use a light path of 5.0 cm for the 
spectrophotometer. 



Rapid Perchloric Acid Digestion for Analysis by Ion Chromatography (Adler 
(1995)): 

Reagents 

1. 70%HNO 3 

2. 70-72% HCIO4 reagent grade 

3. 30% H 2 2 solution 

Procedures 

Add 200 mg (dry wt.) of the sample to a graduated 50 mL digestion (N.P.N.) tube. 
Add 1.0 mL of each HNO3 and HC10 4 to the tube. Place tube into a 300° C preheated 
aluminum digestion block and digest at boiling point until the HNO3 has boiled off (10 
min). This is indicated by the subsidence of boiling. Then add 1.0 mL of H2O2 to the 
solution and continue digestion for another 20 min. Dilute the solution to 25 mL with 
double deionized water, vortex, and filter solution through a 0.2 mm Gelman ion 
chromatography acrodisc. The sample can then be further diluted for analysis by ion 
chromatography. 



81 



Methods for P Analysis, G.M. Pierzynski (ed) 

Dilution of sample for determining total P depends upon the column setup for ion 
chromatography. Adler (1995) found that a 1:10 dilution of the sample is suitable when 
both Dionex IonPac-AG4A and AS4A columns are used. A 1:50 dilution of the sample 
must be used when only an Dionex IonPac-AG4A column is used. The eluent for either 
column setup should be 1.80 mM Na 2 C0 3 and 1.70 mM NaHC0 3 at a flow rate of 2.0 
mL/min. The regenerant for the suppressor should be 12.5 mm H2SO4 at a flow rate of 3 
mL/min. The sample loop volume should be 50 (iL. Use standards containing equivalent 
concentrations of HCIO4 as digested samples to develop a 3 point standard curve. 

Comments 

Adler (1995) found that the addition of an IonPac-AG4A guard column aided in 
better separation of peaks of PO4 and SO4 in a HCIO4 matrix, and that all ions were 
eluted in less than 10 min. This also allows for up to 75% reduction in run time and the 
use of organic solvents can be avoided. 



References: 

Adler, P.R. 1995. Rapid perchloric acid digestion methods for analysis of phosphorus and 

sulfur in aquacultural waterwater and biosolids by ion chromatography. Commun. 

Soil Sci. Plant Anal. 26:85-90. 
APHA. 1989. Standard Methods for the Examination of Water and Wastewater, 17th 

edition. American Public Health Association, Washington, D.C. 
Dick, W.A., and M.A. Tabatabai. 1977. An alkaline oxidation method for determination 

of total phosphorus in soils. Soil Sci. Soc. Am. J. 41:511-514. 
Harris, W.G., H.D. Wang, and K.R. Reddy. 1994. Dairy manure influence on soil and 

sediment compostion: implications for phosphorus retention. J. Environ. Qual. 

23:1071-1081. 
Olsen, S.R. and L.E. Sommers. 1982. Phosphorus, pp. 403-430. In: A.L. Page, R.H. 

Miller, and D.R. Keeney (eds.), Methods of Soil Analysis. 2nd ed. Agronomy Series 

No. 9, Part 2. SoilScience Society of America, Inc., Madison, WI. 
Wen, G., T.E. Bates, R.P. Voroney, J.P. Winter, M.P. Schellenbert. 1997. Comparison of 

phosphorus availability with application of sewage sludge, sludge compost, and 

manure compost. Commun. Soil Sci. Plant Anal. 28:1481-1497. 



82 



Methods for P Analysis, G.M. Pierzynski (ed) 




83 



Methods for P Analysis, G.M. Pierzynski (ed) 

Sample Collection, Handling, Preparation and Storage 

P.M. Haygarth, Institute of Grassland and Environmental Research, England 
A.C. Edwards, Macaulay Land Use Research Institute, Scotland 

Introduction: 

Interfacing between the field, laboratory and chemical analysis is critical in 
determining the forms of phosphorus (P) present in soil- water samples. Collection, 
handling, preparation and storage procedures play a key role the operational definitions 
of P forms (Rowland and Haygarth, 1997) and the lack of a standard protocol can 
introduce serious bias into the precision and accuracy of the determination of the P forms 
(Haygarth et al., 1995). It is therefore essential to adhere to sensible protocols. 

Nomenclature: 

To understand the problems of collection, handling, preparation, and storage it is first 
necessary to consider the definitions and nomenclature of P forms which may be 
determined in a water sample: Collection, handling, preparation, and storage can directly 
affect the analytical endpoint. Forms of P in water attract differing and confusing 
nomenclature, and a systematic and logical means of classification is required. Some 
fraction of the total P content of any water has previously been classified by names which 
define the P in terms of filtration, and subsequently chemical (i.e., Murphy and Riley 
(1962) molybdenum (Mo) blue reaction) methodologies (Haygarth et al., 1998). 
Filtration is strictly a physically based definition of the carrier rather than P form, but has 
been used to define the difference between "soluble" or "dissolved" and "particulate" 
forms. However, any classification of nomenclature based on "dissolved," "soluble," or 
"particulate" is potentially flawed, because (a) different laboratories use different filter 
sizes and (b) P can be associated with a continuum of <0.45 urn sized particles/colloids, 
and samples vary widely in size distribution of particulate/colloidal material (De Haan et 
al, 1984; Haygarth et al, 1997). 

There are similar problems with chemically based definitions. Traditionally the 
Murphy- Riley method has been the standard, but this has been subject to many 
modifications, and there are also uncertainties about what forms of P are determined. 
More recently, ion chromatography and inductively coupled plasma techniques have 
become more popular, but these determine different forms of P than the Murphy- Riley 
reaction. Users therefore need to be aware that P forms are very much methodology 
defined, and the problem is identifying exactly what P forms are determined by each 
method, and ultimately finding a system of nomenclature to incorporate these difficulties. 
Because of this, methodology definitions should be used in the nomenclature where 
possible. 

Reactive P is defined as that which is readily determined analytically by the Mo blue 
reaction (Murphy and Riley, 1962). This is a very specific color reaction that determines 
orthophosphate, but the conditions prior to determining the blue color can change the 
composition of the sample. This means that Mo blue methodology is prone to 
overestimating P, in comparison to chromatographic determinations (Denison et al., 
1998; Edwards and Withers, 1998), because the procedure may also determine loosely 



84 



Methods for P Analysis, G.M. Pierzynski (ed) 

bound inorganic/organic forms of P, by either acid-enhanced hydrolysis (Tarapchak, 
1993) or hydrous ferric oxide-orthophosphate. The reaction is also vulnerable to 
interferences with silica (Ciavatta et al., 1990). Conversely, a sample that requires 
digestion prior to analysis should be called unreactive P. Unreactive P will contain 
organic forms and some condensed forms of P, such as polyphosphates (Ron Vaz et al., 
1993). Therefore any attempt to classify P as "orthophosphate," "organic," or 
"inorganic" in context with Murphy-Riley chemistry is technically incorrect, as it relies 
on the Mo-reaction. Methodology defined terms for describing the P chemistry with the 
Murphy-Riley reaction are therefore "reactive P"(RP), "unreactive P"(UP) and "total 
P"(TP) (i.e., reactive + unreactive, occurring after an appropriate method of digestion, or 
measured directly in an Inductively Coupled Plasma (ICP) system). Thus RP, UP or TP 
are the three prefixes of the suggested nomenclature. 

Similarly, a systematic nomenclature for filtration is proposed, to be used as a suffix 
after chemical form, which removes ambiguity associated with terms like "soluble," 
"dissolved" or "particulate," all of which are non-exacting and subjective. Samples are 
defined specifically according to filter size, with a suffix denoting the pore size (in 
microns) of the filter used in parenthesis (e.g., <0.45 or >0.45). Therefore the established 
system of classifying dissolved reactive P (DRP) would be replaced by RP(<0.45). 
Where a sample was not subjected to filtration, the suffix (unf) is used. Figure 1 provides 
a visual summary of this nomenclature. Ultimately, researchers can expand and adapt 
this methodology-defined nomenclature to include other analytical methods such as ICP 
or ion chromatography. 

Background: 

An idealized and all-encompassing methodology for sample collection is impossible to 
prescribe because it depends on circumstances and samples. Sampling designs must be 
systematic, defensible and hypothesis-driven and therefore random and non-orthogonal 
sample collection programs are not advisable. Types of soil water samples may vary 
from (1) soil extracts determined in some type of laboratory batch procedure, (2) suction 
cup samples that draw water under tension and, (3) flowing or standing waters. Soil 
extractions are considered in other chapters, but it is necessary to be aware that storing 
and sieving soil samples has been found to have a marked affect on resulting extractable 
soil P characteristics (Chapman et al., 1997). Suction cup samplers present uncertainties 
because they draw water under tension, which may not be representative of "mobile" soil 
water. Users of these techniques need to be aware of these limitations. When sampling 
flowing waters from soils, there are three types of procedures: grab samples, flow 
proportional samples and continuous (regular) samples (Haygarth et al., 1998; Lennox et 
al., 1997). Flow proportional or continuous regular sampling provides a truer estimate if 
determining export coefficients is the aim, whereas grab samples can be used for 
comparative studies of spatial differences at one time. 

Phosphorus is vulnerable to transformations during handling and storage, and there 
have been many publications suggesting recommended handling strategies (Annett and 
Dltri, 1973; Bull et al., 1994; Gilmartin, 1967; Haygarth et al., 1995; Henriksen, 1969; 
Heron, 1962; Krawczyk, 1975; Mackereth et al., 1989). Changes can occur in the long 



85 



Methods for P Analysis, G.M. Pierzynski (ed) 

term (Bull et al., 1994) and short term (Haygarth et al., 1995) and can be classified into 
two types: removal or transformation. 

Removal (or "apparent" removal) occurs by sorption to vessel wall (Latterell et al., 
1974) or precipitation. All forms of P can potentially suffer from removal by 
sorption/precipitation reactions (indirectly affected by pH, redox, DOC, Ca, Al and Fe 
content) with container walls. Colloid and particulate content of the water will also 
provide surfaces for sinks and sources of P. Storage vessel size and material are critical 
at regulating the extent of removal by sorption (Annett and D'ltri, 1973; Haygarth et al., 
1995). Freezing of samples is known to reduce losses by sorption, but is not 
recommended because it causes transformations to occur (Johnson et al., 1975). 

Transformation occurs by either chemical or biological mechanisms (Fitzgerald and 
Faust, 1967; Heron, 1962). The sensitivity to transformations in storage brought about by 
microbial mineralization/immobilization, hydrolysis and cell lysis generally increase with 
the complexity of analysis and fractionation performed. For example, analysis for total P 
will only be vulnerable to sorption/desorption interactions (see Figure 1) with vessel 
walls whereas filtered, Mo reactive and unreactive forms of P are also vulnerable to 
transformations and therefore may require a more stringent sample treatment. The 
presence/absence of chemical or biological preservative has been shown to affect 
transformations and Krawczyk (1975) demonstrated that HgCl 2 at an equivalent 
concentration of 400 mg/L suppressed microbial transformations, but has the 
disadvantage of suppressing Mo-blue color reaction in flow-injection systems (Haygarth 
et al., 1995). Other preservatives, such as chloroform, iodine and weak H2SO4 solutions 
have been described (Chakrabarti et al., 1978; Fishman et al., 1986; Murphy and Riley, 
1959), but these techniques can (1) kill microbial populations - releasing reactive P, and 
(2) hydrolyse organic/polyphosphate P. Removal of light and reduction of temperature 
has been shown to have a direct effect on transformations (Haygarth et al., 1995). 
Freezing as a method of preventing transformation is not advisable because it ruptures 
cells and releases P from microorganisms into the soluble phase (Nelson and Romkens, 
1972). 

Sizes and types of filters affect the concentrations of P determined (Haygarth et al., 
1997) and, although threshold sizes used vary from 0.2 to 0.5 urn, 0.45 urn cellulose- 
nitrate- acetate (CNA) filters are most common. Pressure of filtration affects the gas 
propensity for particle retention. The relationship between soluble and particulate P is 
not fixed, but depends on the subsequent storage time and conditions. Samples with a 
high particulate content may tend to block filters during filtration. 

Recommended 'Best Practice': 

Since the range and permutations of sample type, experimental conditions and 
requirements are very high, we are reluctant to recommend a stringent "best procedure." 
One of the key conclusions of Haygarth et al. (1995) was that the range of conditions and 
recommendations by researchers vary in response to different types of sample. For 
example, a suction cup sample from a chalk soil with a high Ca content may require a 
different set of storage conditions than a sample of drainage water from plots recently 
treated with cattle slurry: The former may be vulnerable to removal by Ca-P 
precipitation, while the sample influenced by slurry may be more vulnerable to microbial 
transformations. Further, since the kinetics of change during storage are extremely 



86 



Methods for P Analysis, G.M. Pierzynski (ed) 



PHOSPHORUS IN WATER SAMPLE 



Filtration thorugh 0.45 \xm 
CNA membrane filters 



Appropriate method of digestion 
(NB. Techniques such as ICP) 



Molybdate-blue Reaction (after Murphy and Riley, 1962) 



Reactive P 

unfiltered 

RP . 

unl 



Reactive P 

filtered 
(<0.45 iim) 

ft n 

<0.45jlm 



Total P 


filtered 


(<0.45 iim) 


TP 

< 0.4 5 Jim 



Total P 

unfiltered 

TP . 

unr 



Calculation by difference 



RP. 



Reactive P (>0.45) = RP„ n[ -~ <0 . 4Smm 

Unreactive P «0.45) = TP <MlBB - RP <MJ „ 

Total P (>0.45) = TP „ n( - TP <0 

Unreactive P 00.45) = Total P 00.45) - Reactive POO. 45) 



Figure 1. Operationally defined forms of P in water samples. 
CNA = cellulose-nitrate-acetate, ICP=inductively coupled 
plasma. 



87 



Methods for P Analysis, G.M. Pierzynski (ed) 

variable between water samples, uniform procedures may not help as a broad-brush 
recommendation for all samples. On this cautionary note, we therefore recommend a 
"best practice" rather than a "best procedure." Researchers must be aware of the 
potential hazards and be ready to adapt the procedures to suit their particular 
circumstances. 

Equipment: 

With field sampling, ceramic suction cups may have a tendency to sorb P, whereas 
PTFE samplers may present less of a problem. Storage vessels made with PTFE may 
minimize sorption, but the removal effects of using polyethylene are only slightly worse 
(Haygarth et al., 1995). Sample bottles should be as large as is practicable because this 
reduces the surface area to volume ratio, with volumes >100 mL most effective for 
minimizing changes. Researchers need to consider whether bottles should be washed 
(e.g., 10% v/v H2SO4 or in a P free detergent such as Decon) and if so, how often and the 
appropriate rinsing procedure. If bottles are to be used again perhaps it may be more 
appropriate to store them in clean water. Filtration usually should be through 0.45-um 
CNA membranes, according to the water industry standards. 



Reagents: 

No chemical preservatives should be used, as these change microbial populations, 
which affect the forms of P determined. In extreme circumstances, with waters that are 
particularly vulnerable to transformations, researchers may wish to consider the relative 
advantage of using a 0.22-um CNA membrane to sterilize by filtration. 



Procedure: 

1. When sampling, three bottle fills should be discarded and the fourth sample 
retained, in order to "condition" the bottle. This may be difficult to achieve with 
an autosampler. 

2. Samples must be rapidly transferred to the laboratory and stored in a refrigerator at 
4°C. 

3. The pressure of filtration should not ordinarily exceed 60 cm /Hg (80 kPa). All 
filtration should be undertaken within 12 h. Filters should be pre-washed with 
deionized water, conditioned with sample, and both these eluent solutions 
discarded. 

4. For samples that are vulnerable to transformation (such as those for 
reactive/unreactive P), the total time between sampling and analytical 
determination should not be greater than 24 h. Researchers should be aware of the 
potential for transformations to occur when samplers store storm samples at 
remote sites. 

5. For samples only vulnerable to removal (such as those for total P determination), 
the total time between sampling and analysis can be longer than 24 h, most ideally 
stored at 4°C. It is recommended that if samples must be stored for TP, where 
digestions are needed, they should be readily pipetted into bottles for digestion 
prior to storage to minimize problems of sorption to bottle walls. 



88 



Methods for P Analysis, G.M. Pierzynski (ed) 

Comments: 

The wide range of sample properties means that it is difficult to set a standard 
protocol and the above recommendations must be interpreted in this context. For 
example, there may be a need for setting different protocols for different extremes of 
particulate content or electrolyte concentration. The main principle behind sampling is 
minimal disturbance and rapid transfer to the analytical end point. There is a need to be 
aware of the varying methodological definitions of P, controlled by the analytical 
methods. Recognize that storage starts in the field - perhaps in the suction cup collection 
vessel or in an autosampler bottle, so this must be borne in mind in adopting a best 
practice. Quality control and quality assurance schemes, which use real samples, are to 
be encouraged and adopted. 

References: 

Annett, C.S., and F.M. D'ltri. 1973. Preservation of total soluble phosphorus. In 

"Proceedings 16th Conference Great Lake Research," pp. 214-220. International 

Association Great Lakes Research. 
Bull, K. R., K.H. Lakhani, and A.P. Rowland. 1994. Effects of chemical preservative and 

temperature storage conditions on cations and anions in natural water. Chemistry and 

Ecology 9: 47-62. 
Chakrabarti, C. L, K.S. Subramanian, J.E. Sueiras, and D.J. Young. 1978. Presevation of 

some anionic species in natural waters. J. Amer. Water Works Assoc. October 1978, 

560-565. 
Chapman, P.J., C.A. Shand, A.C. Edwards, and S. Smith. 1997. Effect of storage and 

seiving on the phosphorus composition of soil solution. Soil Sci. Soc. Am. J. 61:315- 

321. 
Ciavatta, C, L.V. Antisari, and P. Sequi. 1990. Interference of soluble silica in the 

determination of orthophosphate-phosphorus. J. Environ. Qual. 19:761-764. 
De Haan, H., T. De Boer, J. Voerman, A.J. Kramer, and J.R. Moed. 1984. Size classes of 

"dissolved" nutrients in shallow, alkaline, humic and eutrophic Tjeukemeer, The 

Netherlands, as fractionated by ultrafiltration. Verh. Int. Ver., Theor. Angew. Limnol. 

22:876-881. 
Denison, F.H., P.M. Haygarth, W.A. House, and A.W. Bristow. 1998. The measurement 

of dissolved phosphorus compounds: Evidence for hydrolysis during storage and 

implications for analytical definitions in environmental analysis. International Journal 

of Environmental Analytical Chemistry. 69: 1 1 1- 123. 
Edwards, A.C, and P.J.A. Withers. 1998. Soil phosphorus management and water 

quality: A UK perspective. Soil Use Manage. 14:124-130. 
Fishman, M.J., L.J. Schroder, and M.W. Shockey. 1986. Evaluation of methods for 

preservation of water samples for nutrient analysis. International Journal of 

Environmental Studies 26:231-238. 
Fitzgerald, G.P., and S.L. Faust. 1967. Effect of water sample preservation methods on 

the release of phosphorus from algae. Limnol. and Oceanog. 12:332-334. 
Gilmartin, M. 1967. Changes in inorganic phosphate concentration occurring during 

seawater sample storage. Limnol. and Oceanog. 12:325-328. 
Haygarth, P.M., CD. Ashby, and S.C Jarvis. 1995. Short term changes in the molybdate 

reactive phosphorus of stored soil waters. J. Environ. Qual. 24:1133-1140. 



89 



Methods for P Analysis, G.M. Pierzynski (ed) 

Haygarth, P.M., L. Hepworth, and S.C. Jarvis. 1998. Forms of phosphorus transfer in 

hydrological pathways from soil under grazed grassland. European J. Soil Science 

49:65-72. 
Haygarth, P. M., M.S. Warwick, and W.A. House. 1997. Size distribution of colloidal 

molybdate reactive phosphorus in river waters and soil solution. Water Res. 31:439- 

442. 
Henriksen, A. 1969. Preservation of water samples for phosphorus and nitrogen 

determination. Vatten 25:247-254. 
Heron, J. 1962. Determination of phosphate in water after storage in polyethylene. 

Limnol. and Oceanog. 7:316-321. 
Johnson, A.H., D.R. Bouldin, and G.W. Hergert. 1975. Some obsevations concerning 

preparation and storage of stream samples for dissolved inorganic phosphate analysis. 

Water Resourc. Res. 11:559-562. 
Krawczyk, D.F. 1975. Preservation of wastewater effluent samples for forms of nitrogen 

and phosphorus. Philadelphia, 152-163. In "Water Quality Parameters," pp. 152-163. 

American Society for Testing and Materials. 
Latterell, J.J., D.R. Timmons, R.F. Holt, and E.M. Sherstad. 1974. Sorption of 

orthophosphate on the surface of water sample containers. Water Resourc. Res. 

10:865-889. 
Lennox, S.D., R.H. Foy, R.V. Smith, and C. Jordan. 1997. Estimating the contribution 

from agriculture to the phosphorus load in surface water. In "Phosphorus Loss from 

Soil to Water" (H. Tunney, O.T. Carton, P.C. Brookes and A.E. Johnston, eds.). CAB 

International, Oxford. 
Mackereth, F.J.H., J. Heron, and J.F. Tailing. 1989. "Water Analysis," Titus Wilson and 

Sons Ltd., Kendal. 
Murphy, J., and J.P. Riley. 1959. The storage of sea-water samples for the determination 

of dissolved inorganic phosphate. Anal. Chim. Acta 14:318-319. 
Murphy, J., and J.P. Riley. 1962. A modified single solution method for the 

determination of phosphate in natural waters. Anal. Chim. Acta 27:31-36. 
Nelson, D.W., and M.J.M. Romkens. 1972. Suitability of freezing as a method of 

preserving runoff samples for analysis of soluble phosphate. J. Environ. Qual. 1:323- 

324. 
Ron Vaz, M.D., A.C. Edwards, CA. Shand, and M.S. Cresser. 1993. Phosphorus 

fractions in soil solution: Influence of soil acidity and fertilizer additions. Plant Soil 

148:179-183. 
Rowland, A. P., and P.M. Haygarth. 1997. Determination of total dissolved phosphorus in 

soil solutions. J. Environ. Qual. 26: 410-415. 
Tarapchak, S.J. 1993. Soluble reactive phosphorus in lake water: Evidence for 

molybdate enhanced hydrolysis. J. Environ. Qual. 12:105-108. 



90 



Methods for P Analysis, G.M. Pierzynski (ed) 

Analyzing for Dissolved Reactive Phosphorus in Water 
Samples 

D.H. Pote, USDA-ARS, Booneville, AR 
T.C. Daniel, University of Arkansas 

Introduction: 

Dissolved reactive P (DRP), sometimes called soluble reactive P, refers to the P 
fraction that passes through a 0.45-um-pore-diameter membrane filter and responds to 
the molybdate colorimetric test without preliminary hydrolysis or oxidative digestion of 
the water sample. It is largely a measure of dissolved orthophosphate, the form of P most 
readily available to aquatic plants, and thus is often considered the most critical P fraction 
contributing to accelerated eutrophication of surface waters. Although filtration through 
a 0.45-um pore diameter membrane filter may not completely separate dissolved and 
suspended forms of P, this method can be easily replicated. Therefore, it provides a 
convenient technique for clearly defining the analytical separation of the dissolved and 
suspended P fractions. 

Development of the molybdate colorimetric test for ortho-P in water samples was 
based on the observation that ammonium molybdate and potassium antimony tartrate 
react with dilute ortho-P solutions in an acid medium to form an antimony-phospho- 
molybdate complex. Reduction of this complex by ascorbic acid gives it an intense blue 
color that is proportional to the ortho-P concentration. Early prototypes of this 
colorimetric technique have been used for more than 60 years to determine P 
concentrations. Ammon and Hinsberg (1936) reported using ascorbic acid to reduce 
phosphomolybdic acid to molybdenum blue as a method of analyzing for P and As. 
Greenfield and Kalber (1954) suggested using the technique for analysis of sea water. 
Murphy and Riley (1958) recommended altering the method to provide a single reagent 
for phosphate determination in sea water, but their initial modified technique required 24 
h at room temperature or 30 min at 60°C for full color development. The higher 
temperature or long time period required for color development raised concerns because 
either condition may allow hydrolysis of some organic P compounds to orthophosphate. 
Therefore, Murphy and Riley (1962) revised the method again when they found that 
adding antimony (as potassium antimonyl tartrate) to the reagent caused full color 
development in 10 min at room temperature. The basic procedure has changed little 
since 1962, but it has been modified for use on autoanalyzers. 

For the procedure as described below, the minimum detectable P concentration is 
approximately 10 ug/L. 

Equipment: 

1. Filtration apparatus (0.45-um pore diameter) 

2. Photometer - Spectrophotometer with infrared phototube for use at 880 nm and 
providing a light path of at least 2.5 cm or a filter photometer with a red color filter 
and a light path of at least 0.5 cm. For light path lengths of 0.5, 1.0, and 5.0 cm, 
the P ranges are 0.3-2.0, 0.15-1.30, and 0.01-0.25 mg/L, respectively. 



91 



Methods for P Analysis, G.M. Pierzynski (ed) 

3. Acid-washed glassware and plastic bottles: graduated cylinders (5 mL to 100 mL 
measurements), volumetric flasks (100 mL, 500 mL, and 1000 mL), storage bottles 
(including dark glass-stoppered, and opaque plastic), pipets, eye droppers, and test 
tubes or flasks for reading sample absorbance 



Reagents: 

1 . 2.5 M H 2 S0 4 . Slowly add 70 mL of concentrated H 2 S0 4 to approximately 400 mL 
of distilled water in a 500 mL volumetric flask. After the solution has cooled, 
dilute to 500 mL with distilled water, mix, and transfer to a plastic bottle for 
storage. 

2. Ammonium molybdate solution. Dissolve 20 g of (NH 4 ) 6 Mo7 24 ' 4H 2 in 500 
mL of distilled water. Store in a plastic bottle at 4°C. 

3. Ascorbic acid, 0. 1 M. Dissolve 1 .76 g of ascorbic acid in 100 mL of distilled 
water. The solution is stable for about a week if stored in an opaque plastic bottle 
at4°C. 

4. Potassium antimonyl tartrate solution. Using a 500 mL volumetric flask, dissolve 
1.3715 g of K(SbO)C 4 H 4 6 1/2 H 2 in approximately 400 mL of distilled water, 
and dilute to volume. Store in a dark, glass- stoppered bottle. 

5. Combined reagent. When making the combined reagent, all reagents must be 
allowed to reach room temperature before they are mixed, and they must be mixed 
in the following order. To make 100 mL of the combined reagent: 

a. Transfer 50 mL of 2.5 M H 2 S0 4 to a plastic bottle. 

b. Add 15 mL of ammonium molybdate solution to the bottle and mix. 

c. Add 30 mL of ascorbic acid solution to the bottle and mix. 

d. Add 5 mL of potassium antimonyl tartrate solution to the bottle and mix. If 
turbidity has formed in the combined reagent, shake and let stand for a few 
min until turbidity disappears before proceeding. Store in an opaque plastic 
bottle. The combined reagent is stable for less than 8 h, so it must be freshly 
prepared for each run. 

6. Stock phosphate solution. Using a 1000 mL volumetric flask, dissolve 219.5 mg 
anhydrous KH 2 P0 4 in distilled water and dilute to 1000 mL volume; 1 mL 
contains 50 ug of P. 

7. Standard P solutions. Prepare a series of at least six standard P solutions within the 
desired P range by diluting stock phosphate solution with distilled water. 

8. Phenolphthalein indicator solution. 

Procedure: 

1. Filter sample through a membrane filter (0.45-um pore diameter). Hard-to-filter 
samples can be prefiltered through a glass fiber filter to prepare them for 
membrane filtration. 

2. Pipet 50.0 mL of sample into a clean, dry test tube or flask. Add 1 drop (0.05 mL) 
of phenolphthalein indicator and mix. If a red color develops, add just enough 
drops of 2.5 M H 2 S0 4 to remove the color. Add 8.0 mL of combined reagent and 
mix thoroughly. Wait at least 10 min (but no more than 30 min) before measuring 



92 



Methods for P Analysis, G.M. Pierzynski (ed) 

the absorbance of each sample at 880 nm, using reagent blank as the reference 
solution. 

3. Natural color of water should not interfere at the high wavelength used in this 
procedure. However, if the water samples are turbid or strongly colored, prepare a 
blank by adding all reagents except potassium antimonyl tartrate and ascorbic acid 
to a water sample. To obtain the actual absorbance of each sample, subtract 
absorbance of the blank from the sample's measured absorbance. 

4. Prepare a calibration curve from the series of at least six standard P solutions 
within the desired P range. Use a distilled water blank with the combined reagent 
when making the photometric readings for a calibration curve, and plot absorbance 
vs. P concentration to obtain a straight line passing through the origin. Each set of 
samples should include at least one P standard to assure accuracy of the results. 

Comments: 

Arsenate concentrations as low as 0.1 mg/L can interfere with the P determination by 
reacting with the molybdate reagent to produce a blue color. Hexavalent chromium and 
N0 2 " at 1 mg/L can interfere to give results about 3% low, and at 10 mg/L give results 
10-15% low. 

If an autoanalyzer is being used for this procedure, the following adjustment in reagent 
preparation is recommended: When making potassium antimonyl tartrate solution, 1.5 g 
of K(SbO)C 4 H 4 6 1/2H 2 should be dissolved in distilled water to make 500 mL of 
solution. 



References: 

Amnion, R., and K. Hinsberg. 1936. Colorimetric determination of phosphorus and 

arsenic with ascorbic acid. Hoppe-Seyl. Z. 239:207. 
Greenfield, L.J., and F.A. Kalber. 1954. Inorganic phosphate measurement in sea water. 

Bull. Marine Sci. Gulf Caribbean 4:323-335. 
Murphy, J., and J. P. Riley. 1958. A single- solution method for the determination of 

soluble phosphate in natural waters. J. Marine Biol. Assoc. UK 37: 9. 
Murphy, J., and J. P. Riley. 1962. A modified single solution method for the 

determination of phosphate in natural waters. Anal. Chim. Acta 27:31-36. 



93 



Methods for P Analysis, G.M. Pierzynski (ed) 

Analyzing for Total Phosphorus and Total Dissolved 
Phosphorus in Water Samples 

D.H. Pote, USDA-ARS, Booneville, AR 
T.C. Daniel, University of Arkansas 

Introduction: 

Dissolved orthophosphate is the form of P most readily available to aquatic plants, but 
numerous studies have shown that other forms of P can be hydrolyzed to the 
orthophosphate form in wastewater-treatment facilities and in natural waters. Therefore, 
when assessing the long-term potential for accelerated eutrophication of surface water 
due to P loading, many researchers and watershed managers want to know the total P 
concentration (regardless of P form) in water samples. 

Polyphosphates and phosphates bound to organic substances do not react with the 
molybdate reagent used for colorimetric P analysis. Therefore, analysis for total P 
content of water samples requires that all condensed and organic P compounds, including 
particulate P, first be converted (hydrolyzed) to orthophosphate so they can be 
determined colorimetrically. This is accomplished by digesting the sample in strong acid 
at high temperature to oxidize the organic matter and release P as orthophosphate. 
Published methods for accomplishing the digestion process have been available for many 
decades. Improved methods have been developed, but all of them use heat and/or various 
strong acids, sometimes in combination with strong oxidizing reagents. For example, the 
wet ashing digestion method (using concentrated HNO3 and H2SO4) described by Peters 
and Van Slyke (1932) was considered reliable, but was very time-consuming, so other 
researchers developed faster digestion procedures. Perchloric acid digestion, described 
by Robinson (1941), is still considered a standard method for total P analysis, but it is 
time-consuming and dangerous because heated mixtures of HCIO4 and organic matter 
may explode violently. Therefore, other digestion methods (listed below) are usually 
preferred. 

To determine the total dissolved P fraction, the particulate P is separated by filtering 
the water sample through a 0.45 urn pore diameter membrane filter before beginning the 
digestion procedure. To determine total P (dissolved + particulate), an unfiltered sample 
is shaken (to suspend the particulate matter) just before measuring the subsample for 
digestion. 

Sulfuric Acid - Nitric Acid Digestion Method: 

Equipment 

1. Digestion rack. Digestion racks designed for micro-Kjeldahl digestions can be 
used, but need to include a provision for withdrawal of fumes. A digestion rack 
heated by either gas or electricity is suitable. 

2. Micro-Kjeldahl flasks. 

3. Acid- washed graduated cylinders, pipets, eye droppers, and 100 mL volumetric 
flasks. 

4. Any additional equipment required for colorimetric determination of P in the 
digested sample solution (described in the Dissolved Reactive P section). 



94 



Methods for P Analysis, G.M. Pierzynski (ed) 

Reagents 

1. Concentrated H2SO4 

2. Concentrated HNO3 

3. Phenolphthalein indicator aqueous solution 

4. lMNaOH 

5. Any additional reagents required for colorimetric determination of P in the 
digested sample solution (described in the Dissolved Reactive P section) 

Procedure 

1. Transfer a measured volume of sample into a micro-Kjeldahl flask. We 
recommend a volume of at least 25 mL if adequate sample is available. Larger 
volumes can be used, but they require a longer digestion time. 

2. Add 1 mL of concentrated H 2 SO4 

3. Add 5 mL of concentrated HNO3 

4. Digest to a volume of 1 mL and then continue digesting until the solution becomes 
colorless (to remove the HNO3 ) 

5. Cool the flask and add approximately 20 mL of distilled water. 

6. Add 1 drop (0.05 mL) of phenolphthalein indicator and mix. 

7. Add drops of 1 M NaOH until the sample solution acquires a faint pink tinge. 

8. Transfer the neutralized solution (if necessary, filtering to remove turbidity or 
particles) into a 100-mL volumetric flask. If a filter is used, be sure to add 
distilled- water filter washings to the flask. 

9. Adjust sample volume to 100 mL with distilled water. 

10. Use the molybdate colorimetric test (described in previous chapter on Dissolved 
Reactive P) to determine the P content of the digested solution. 

11. To prepare the calibration curve, carry a series of standards through the digestion 
process. Do not use standards that have not been digested. 



Persulfate Digestion Method: 

Equipment 

1. Hot plate with adequate heating surface. An autoclave or pressure cooker capable 
of developing 98 - 137 kPa may be used instead of a hot plate. 

2. Acid- washed graduated cylinders, pipets, eye droppers, and volumetric flasks (100 
mL and 1000 mL). 

3. Any additional equipment required for colorimetric determination of P in the 
digested sample solution (described in the Dissolved Reactive P section). 

Reagents 

1 . Phenolphthalein indicator solution 

2. Sulfuric acid solution. Transfer approximately 600 mL of distilled water to a 1000 
mL volumetric flask. Slowly (and carefully) add 300 mL of concentrated H 2 SO4 . 
After the solution has cooled, dilute to 1000 mL with distilled water and mix. 

3. Ammonium persulfate, (NH 4 ) 2 S2 08 solid or potassium persulfate, K2S2O8 solid. 

4. lMNaOH 

5. Any additional reagents required for colorimetric determination of P in the 
digested sample solution (described in the Dissolved Reactive P section) 



95 



Methods for P Analysis, G.M. Pierzynski (ed) 

Procedure 

1. Thoroughly mix the sample, and measure a suitable portion (50 mL is 
recommended) into a flask. 

2. Add 1 drop (0.05 mL) of phenolphthalein indicator and mix. If a red color 
develops, add just enough drops of H2SO4 to remove the color. 

3. Add 1 mLof H2SO4 solution. 

4. Add either 0.4 g of solid (NFL^Og or 0.5 g of solid K 2 S 2 8 and mix. 

5. Boil the sample solution gently on the preheated hot plate for at least 30-40 min or 
until the volume is reduced to 10 mL. Some organophosphorus compounds may 
require 2 h for complete digestion. 

6. Cool the solution, and dilute to approximately 30 mL with distilled water. 

7. Add 1 drop (0.05 mL) of phenolphthalein indicator. 

8. Add drops of 1M NaOH until the sample solution is neutralized (acquires a faint 
pink tinge). 

9. Dilute to 100 mL volume with distilled water. If a precipitate forms, do not filter, 
but shake well for any subdividing of the sample. The precipitate redissolves 
during the colorimetric test due to increased acidity. 

10. Use the molybdate colorimetric test (described in previous chapter on Dissolved 
Reactive P) to determine the P content of the digested solution. 

11. To prepare the calibration curve, carry a series of standards through the digestion 
process. Do not use standards that have not been digested. 



Kjeldahl Digestion Method: 

The Kjeldahl digestion procedure also converts condensed and organic P compounds, 
including particulate P, to orthophosphate. Therefore, if the water samples are being 
digested by the Kjeldahl method to determine their total Kjeldahl nitrogen content, then 
total P can also be measured (without further digestion) by simply using the molybdate 
colorimetric test (described in the previous chapter on Dissolved Reactive P) to determine 
the P content of the digested solution. To prepare the calibration curve, carry a series of 
standards through the Kjeldahl digestion process. 

Calculations: 

For any of the three digestion methods listed above, always use the correct dilution 
ratio when calculating the total P concentration in the original sample. For example, if a 
50-mL sample is used, and the sample is diluted to a final volume of 100 mL following 
the digestion procedure, then the measured concentration should be multiplied by 2 to 
obtain the concentration in the original water sample. 

TotalDilutedVolume(mL) 
Total P (mg / L) = P concentration in analyzed solution (mg / L) x 



OriginalSampleVolume(mL) 

Comments: 

The sulfuric acid - nitric acid digestion method is recommended for most samples. 
The persulfate digestion method is much simpler to use and usually gives excellent 



96 



Methods for P Analysis, G.M. Pierzynski (ed) 



recovery rates, but when digesting potentially difficult samples, it should probably be 
checked against the sulfuric acid - nitric acid digestion and adopted if identical recoveries 
are obtained. 



References: 

Peters, J.P., and D.D. Van Slyke. 1932. Quantitative Clinical Chemistry. Vol II. 

Methods. The Williams and Wilkins Co., Baltimore, MD. 
Robinson, R.J. 1941. Perchloric acid oxidation of organic phosphorus in lake waters. 

Ind. Eng. Chem. Anal. (Ed.) 13:465-466. 



97 



Methods for P Analysis, G.M. Pierzynski (ed) 

Using the Iron Oxide Method to Estimate Bioavailable 
Phosphorus in Runoff 

R.G. Myers, Kansas State University 
G.M. Pierzynski, Kansas State University 

Introduction: 

The use of iron-oxide (FeO) coated paper to test soil was first reported by Sissingh 
(1983), who wanted to develop a soil phosphorus (P) test that would estimate plant- 
available P in tropical soils without mobilizing other forms of phosphates. A strip of 
filter paper impregnated with iron hydroxide functioned as a P sink and adsorbed mobile 
P from solution, so Sissingh (1983) called the analyzed P, the Pi value (i referring to iron 
hydroxide). Interest in the method was soon extended to a wider range of soils (Menon et 
al., 1989). The test has an advantage over standard soil P tests because the FeO paper 
functions as an ion sink and doesn't react with soil as do chemical extractants. A unique 
feature of the FeO method rests in its inherent preferential selectivity of FeO for P ions 
over all other anions found in soil, except OH (Menon, 1993; van der Zee et al., 1987). 

The FeO test has been identified by quite a number of different terms in various papers 
and publications, e.g., Pi test, Fe-oxide strip method, and Pi test (Chardon et al., 1997; 
Perrot and Wise, 1993; Sharpley, 1993a). To avoid confusion, P extraction by FeO- 
coated paper will be called the FeO method, and the P extracted will be called FeO-P. 

Interest in applying the FeO method to agricultural runoff has been developing 
recently in an effort to assess the potential of P in runoff to stimulate freshwater 
eutrophication. The bioavailable P content (BAP) of dilute runoff sediment assessed by 
the FeO method was related (r 2 = 0.63-0.96) to the growth of P-starved algae 
(Selanastrum capricornutum) (Sharpley, 1993a). Additional work showed that FeO-P 
from runoff sediment was related (P > 0.001) to algal growth in Anabaena, 
Ankistrodesmus, and Euglena (Sharpley, 1993b). The FeO method has the unique 
capability of differentiating soluble inorganic P from FeO-P in sediment of runoff. The 
sediment FeO-P is called bioavailable particulate P (BPP) and is calculated according to 

BPP = total BAP - SP [1] 

where total BAP is total FeO-P from unfiltered runoff, and SP is soluble inorganic P in 
filtered runoff (0.45-um filter). 

The FeO method has a stronger theoretical justification for estimating P availability of 
soil and runoff for plants and algae than do chemical methods (Sharpley, 1993a). The 
rationale for this theoretical justification lies in the mechanism of P adsorption onto the 
FeO-coated paper. Such adsorption closely simulates that of plants and algae and thereby 
gives an estimation of BAP, whereas chemical methods may mobilize additional forms of 
P which are not available to plants or algae. Therefore, the FeO method is an additional 
tool used to assess the potential for runoff to increase fresh-water eutrophication. 

In the past, filter paper with large pores up to 20 to 25 urn sometimes was used to 
make FeO paper, however, there is less tendency for soil particles to become lodged in 
papers with small pores, e.g. < 5.0 urn, so small-pore paper is now recommended 



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Methods for P Analysis, G.M. Pierzynski (ed) 

(Chardon et al., 1997). Traditionally, filter paper circles with a 15-cm diameter were 
coated with FeO by immersing them first in a FeCl3 solution, then after drying, they were 
immersed in an NH4OH solution (van der Zee, et al. 1987). After drying they were cut 
into strips, often 2x10 cm— from whence came the term strip-P. 

Recently, filter circles with a 5.5 cm diameter have been used to make the FeO papers 
instead of cutting strips from the larger circles (Myers et al., 1995, 1997). The surface 
area of the 5.5-cm circles exceeds that of the traditional 2 x 10-cm strips by about 20%; 
however, the primary reason for using circles instead of strips is to eliminate the need for 
cutting strips. Within a 12 h shaking time, each 5.5-cm FeO circle has adequate 
adsorption capacity to remove 99% of the P in a solution containing 16.1 urn P (Myers et 
al., 1997). van der Zee et al. (1987) reported similar results with adsorption of 18 umol 
P after shaking one 2 x 10-cm strip for 20 h. 

Holding the FeO paper in a fixed orientation during shaking helps to prevent soil 
particles from lodging in the pores of the paper and contaminating it (Myers et al., 1995; 
1997). Although runoff aliquots usually contain much less than 1.0 g of sediment, the 
amount of soil used in soil extraction, stabilization of each FeO paper between 
polyethylene screens is still recommended for analysis of runoff samples, some of which 
can contain substantial quantities of sediment. Holding the screens in a fixed orientation 
during shaking also prevents the FeO papers from sticking to the walls of the shaking 
vessel, as often occurs when the papers are allowed to shake freely in solution. Such 
sticking could reduce adsorption effectiveness of the FeO paper. 

A solution of 0.01 M CaCl2 is used as the shaking matrix for the FeO paper and soil 
because deionized water has the tendency to disperse soil, which may then lodge in the 
pores of the filter paper (Sissingh, 1983). This may lead to errors in P analysis (Myers et 
al., 1995); however, runoff has been extracted by the FeO method without addition of any 
CaCl 2 (Sharpley, 1993a). We have found that FeO-P from runoff made with 0.01 M 
CaCb was the same as that from duplicate runoff samples shaken without CaCb (data 
unpublished), but similar results may not always hold true for every type of runoff in 
every location. The potential for significant contamination of FeO papers by not 
amending the runoff with CaCb during shaking may depend upon the clay content of the 
sediment and the P content of the clay as well as the amount of sediment in the runoff. 

Equipment: 

1. End-over-end shakers have been used for the FeO method (Sissingh, 1983; 
Sharpley, 1993). Reciprocating shakers have also been used (Menon et al., 1989; 
Myers et al, 1997). 

2. 2 L beaker 

3. 11 8-mL wide-mouthed glass bottles 

4. 125-mLErlenmeyer flasks 

5. 50-mL Erlenmeyer flasks 

6. Spectra/Mesh polyethylene screens (925 urn, Spectra/Mesh filters, Fisher Co., St. 
Louis; Fisher cat. no. 08-670-175) 

7. Parafilm 



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Methods for P Analysis, G.M. Pierzynski (ed) 

Reagents: 

1. 0.65MFeCl 3 - 6H 2 O + 0.6MHCl 

2. 2.7MNH 4 OH 

3. O.2MH2SO4 

4. Reagents used for the Murphy and Riley (1962) colorimetric procedure 

Procedure: 

We use hardened 5.5 cm circles of Whatman no. 50 filter paper for making the FeO 
paper (Myers et al., 1997). Briefly, we immerse the papers, one by one, in 0.65 M FeCh 
• 6H2 O containing 50 mL of concentrated HC1 per liter of solution, and leave them in the 
container overnight. Chardon et al. (1997) recommend acidification of the FeCh solution 
if the papers are to be stored, thus we acidify with HC1. After air-drying the papers on a 
rack, they are immersed in 2.7 MNH 4 OH for 30 s and then allowed to drain for 15 s 
before thoroughly rinsing in two containers of clean distilled water. They are placed in a 
bucket of clean water for 1 h to permit dissipation of any residual ammonia. The papers 
are then ready to use immediately or they can be dried for later use. For further details on 
paper preparation, see Myers et al. (1997). 

Polyethylene screens are cut approximately 9 cm in diameter from Spectra/Mesh 
filters. These screens are used to enclose each FeO paper during shaking (Myers et al., 
1997). One FeO paper is placed between two of these screens held together by a plastic 
clamp, making a paper-screen assembly to insert into the shaking bottle. 

We have followed the traditional FeO method for determining BAP in runoff 
(Sharpley, 1993a), except that we use a total shaking volume of 80 mL. We add 50 mL 
of runoff plus 30 mL of deionized water. When 80 mL of solution is shaken in 1 18-mL 
bottles orientated horizontally and end-to-end, the shaking action completely rinses the 
sides and top of the bottles with each excursion of the reciprocating shaker. If shaking 
action is adequate in some other type of shaking vessel, the total volume of solution is 
optional and discretionary. Also, for runoff with low levels of FeO-P, 80 mL of runoff 
may be used without adding any water. 

The FeO paper- screen assembly is inserted, clamp end first, into the bottle containing 
runoff. Cover the bottles tightly with a layer of Parafilm, and then screw the closures on 
tightly to seal. The bottles are shaken on a reciprocating shaker for 16 h at a speed of 125 
to 135 excursions/min. Shaking speed can be increased, if needed, to increase mixing. 

After a 16 h shaking period, we remove the papers from the screens and rinse each 
paper under a stream of deionized water for a few seconds. The papers are coiled and 
placed in the neck of a 125-mL Erlenmeyer flask where they may either be left to dry or 
pushed to the bottom and extracted immediately. Extract the P from the papers by adding 
50 mL of 0.2 MH 2 SO4 to flasks and shaking them 1 h at 100 to 125 excursion/min. An 
aliquot of the H 2 SO4 solution is analyzed for P using the Murphy and Riley (1962) after 
neutralization of acidity. For neutralization, phenolphthalein color indicator gives a clear 
end point in the FeO solution. Duplicate, or triplicate, control FeO papers, without any 
soil or runoff, are also shaken and extracted to correct for any P contained in reagents and 
water. For further details on the FeO procedure described above, see Myers et al. (1995, 
1997). 



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Methods for P Analysis, G.M. Pierzynski (ed) 

Calculations: 

The Murphy and Riley (1962) method of P analysis gives FeO-P in ug P/mL. If data 
for FeO-P are presented in units of ug/L then the appropriate calculations for BAP are: 

Total BAP (ug/L) = [volume of H 2 S0 4 (L) x P in H 2 S0 4 (ug/L )] 4- 
[volume of runoff sample extracted with FeO (L)] 

where total BAP is the total bioavailable P in the runoff, and H 2 SO4 is 50 mL of 0.2 
MH2SO4 used to extract P from each FeO paper. Calculations for bioavailable particulate 
P (BPP), the FeO-P associated with the sediment, are given in Eq. 1 above. 



Comments: 

Algae use only the orthophosphate form of P; however, organic forms of P can 
undergo mineralization and also become available (Correll, 1998). Thus, organic P can 
be considered a latent source of BAP. Some discussion has been focused on methods to 
limit hydrolysis of organic P adsorbed onto FeO paper (Robinson and Sharpley, 1994); 
however, it appears that such adsorption and hydrolysis of organic P is not a problem in 
using the FeO method to estimate BAP because organic P may be justifiably classified as 
latent BAP which may be mineralized at any time and thereby become immediately 
available for algal uptake. 



References: 

Chardon, W.J., R.G. Menon, and S.H. Chien. 1997. Iron oxide impregnated filter paper 

(Pi test): A review of its development and methodological research. Nutr. Cycl. 

Agroecosyst. 46:41-51. 
Correll, D.L. 1998. The role of phosphorus in the eutrophication of receiving waters: A 

review. J. Environ. Qual. 27:261-266. 
Menon, R.G. 1993. The Pi test for evaluating bioavailability of phosphorus, p. 58-67. In 

K.B. Hoddinott and T.A. O'Shay (ed.) Application of agricultural analysis in 

environmental studies. ASTM STP 1162, Philadelphia, PA. 
Menon, R.G., L.L. Hammond, and HA. Sissingh. 1989. Determination of plant- available 

phosphorus by the iron hydroxide-impregnated filter paper (Pi ) soil test. Soil Sci. 

Soc. Am. J. 53:110-115. 
Murphy, J., and J. P. Riley. 1962. A modified single solution method for the 

determination of phosphate in natural waters. Anal. Chim. Acta 27:31-36. 
Myers, R.G., G.M. Pierzynski, and S.J. Thien. 1995. Improving the iron oxide sink 

method for extracting soil phosphorus. Soil Sci. Soc. Am. J. 59:853-857. 
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extracting soil phosphorus: Paper preparation and use. Soil Sci. Soc. Am. J. 61:1400- 

1407. 
Perrott, K.W., and R.G. Wise. 1993. An evaluation of some aspects of the iron oxide- 
impregnated filter paper (Pi) test for available soil phosphorus with New Zealand 

soils. N. Z. J. Agric. Res. 36:157-162. 



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Robinson, J.S., and A.N. Sharpley. 1994. Organic phosphorus effects on sink 

characteristics of iron-oxide-impregnated filter paper. Soil Sci. Soc. Am. J. 58:758- 

761. 
Sharpley, A.N. 1993a. An innovative approach to estimate bioavailable phosphorus in 

agricultural runoff using iron oxide-impregnated paper. J. Environ. Qual. 22:597- 

601. 
Sharpley, A.N. 1993b. Estimating phosphorus in agricultural runoff available to several 

algae using iron-oxide paper strips. J. Environ. Qual. 22:678-680. 
Sissingh, H. A. 1983. Estimation of plant-available phosphates in tropical soils. Anew 

analytical technique. Nota 235. Inst, for Soil Fertility Res., Haren, the Netherlands, 
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